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. 2020 May 9;150(7):1722–1730. doi: 10.1093/jn/nxaa113

Cell-Type–Specific, Ketohexokinase-Dependent Induction by Fructose of Lipogenic Gene Expression in Mouse Small Intestine

Arwa Al-Jawadi 1,✉, Chirag R Patel 2,✉, Reilly J Shiarella 3, Emmanuellie Romelus 4, Madelyn Auvinen 5, Joshua Guardia 6, Sarah C Pearce 7,✉, Kunihiro Kishida 8,✉, Shiyan Yu 9, Nan Gao 10, Ronaldo P Ferraris 11,✉
PMCID: PMC7330472  PMID: 32386219

ABSTRACT

Background

High intakes of fructose are associated with metabolic diseases, including hypertriglyceridemia and intestinal tumor growth. Although small intestinal epithelia consist of many different cell types, express lipogenic genes, and convert dietary fructose to fatty acids, there is no information on the identity of the cell type(s) mediating this conversion and on the effects of fructose on lipogenic gene expression.

Objectives

We hypothesized that fructose regulates the intestinal expression of genes involved in lipid and apolipoprotein synthesis, that regulation depends on the fructose transporter solute carrier family 2 member a5 [Slc2a5 (glucose transporter 5)] and on ketohexokinase (Khk), and that regulation occurs only in enterocytes.

Methods

We compared lipogenic gene expression among different organs from wild-type adult male C57BL mice consuming a standard vivarium nonpurified diet. We then gavaged twice daily for 2.5 d fructose or glucose solutions (15%, 0.3 mL per mouse) into wild-type, Slc2a5-knockout (KO), and Khk-KO mice with free access to the nonpurified diet and determined expression of representative lipogenic genes. Finally, from mice fed the nonpurified diet, we made organoids highly enriched in enterocyte, goblet, Paneth, or stem cells and then incubated them overnight in 10 mM fructose or glucose.

Results

Most lipogenic genes were significantly expressed in the intestine relative to the kidney, liver, lung, and skeletal muscle. In vivo expression of Srebf1, Acaca, Fasn, Scd1, Dgat1, Gk, Apoa4, and Apob mRNA and of Scd1 protein increased (P < 0.05) by 3- to 20-fold in wild-type, but not in Slc2a5-KO and Khk-KO, mice gavaged with fructose. In vitro, Slc2a5- and Khk-dependent, fructose-induced increases, which ranged from 1.5- to 4-fold (P < 0.05), in mRNA concentrations of all these genes were observed only in organoids enriched in enterocytes.

Conclusions

Fructose specifically stimulates expression of mouse small intestinal genes for lipid and apolipoprotein synthesis. Secretory and stem cells seem incapable of transport- and metabolism-dependent lipogenesis, occurring only in absorptive enterocytes.

Keywords: chylomicron, epithelia, lipogenesis, lipids, liver, organoids, stem cells, sugars

Introduction

Increased fructose intake leads to portal vein and systemic fructosemia associated with nonalcoholic fatty liver and other metabolic diseases now at epidemic levels in the United States (1, 2). The small intestine plays a vital role in protecting the liver from fructose toxicity, as luminal fructose in modest concentrations can be metabolized intracellularly by intestinal cells into glucose and other metabolites, reducing the amount of fructose reaching the portal vein and liver (3, 4). After absorption, some fructose is converted by the small intestine to lipids, a process that then increases intestinal lipid secretion into the blood. In humans, labeled fructose in a mixed meal resulted in labeled palmitate incorporated in circulating chylomicrons, indicating small intestinal de novo lipogenesis from fructose (5–7). Dietary fructose also stimulated production of human intestinal triglycerides as well as apolipoprotein B48 (the structural protein for chylomicrons, APOB48, an RNA-edited product of the Apob gene) involved in the synthesis of chylomicrons (5, 8). Fructose feeding of hamsters for 3 wk increased not only enterocyte (ENT) incorporation of [3H]acetate into fatty acids but also secretion of intestinal APOB48 and triglycerides (9). Fructose-fed, nonobese, adenomatous polyposis coli knockout (KO) mice (a model of intestinal tumorigenesis) not only converted fructose to but also increased synthesis of fatty acids that supported tumor growth (10). Because fructose increases the amounts of some inflammatory cytokines that alter the Wnt pathway, it can disrupt epithelial differentiation, leading to precancerous ectopic formation in the villi of ENTs inappropriately expressing stem cell markers (11). Thus, fructose-induced intestinal lipogenesis may contribute to metabolic diseases and also to the development of intestinal and colon cancer.

Because absorptive ENTs constitute ∼70–80% of all small intestinal cells, studies attribute these fructose-induced responses to be solely due to this cell type (12). The adult small intestinal epithelium, however, is composed of intestinal stem cells (ISCs) that continuously proliferate and differentiate into absorptive ENTs as well as secretory enteroendocrine, Paneth (PAN), goblet (GOB), and other cells with unknown capacities for lipid synthesis and secretion.

Fructose is transported across the apical membrane and down its chemical gradient into the cytosol via the intestinal fructose transporter glucose transporter 5 [GLUT5, encoded by solute carrier family 2 member a5 (Slc2a5)] (13, 14). Fructose is then rapidly metabolized by ketohexokinase (KHK, encoded by Khk) and aldolase into 3-carbon intermediates that can be converted to acetyl-CoA acting as precursors for the synthesis of triglycerides (15) in the intestinal cell. Some fructose is transported out of the intestine by GLUT2 (encoded by Slc2a2) into the hepatic portal vein and to the liver, where fructose is more lipogenic than glucose because it regulates the expression levels of sterol regulatory element-binding transcription factor 1 (Srebf1) that modulates expression of downstream lipogenic genes (16, 17) and because it is catabolized faster than glucose (18), thereby rapidly providing precursor carbons for fatty acid synthesis (19). Hepatic and intestinal lipogenesis begins with acetyl-CoA carboxylase 1 (encoded by Acaca) converting acetyl-CoA into malonyl-CoA, which is utilized by fatty acid synthase (encoded by Fasn) to produce palmitic acid. Palmitic and other longer chain SFAs can be desaturated by stearoyl-CoA desaturase 1 (SCD1, encoded by Scd1). During triglyceride synthesis, fatty acids are attached to monoacylglycerol, via monoacylglycerol acyltransferases, to make diacylglycerol and then, via diacylglycerol acyltransferases (DGAT1, encoded by Dgat1), to triglycerides. In the small intestine, DGAT1 is the isoform that synthesizes postprandial triglycerides (20), which are then packaged into chylomicrons coated with intestine-specific apolipoproteins APOB48 (encoded by Apob) and APOA4 (encoded by Apoa4) and other apolipoproteins. Chylomicrons, which are made only in intestinal cells, are then exocytosed to the lymph circulation. The proportion of chylomicrons from fructose-induced lipogenesis relative to total chylomicrons is likely related to dietary fructose concentrations.

Although many previous studies have demonstrated intestinal lipogenesis from fructose, no studies have examined the fructose-induced expression of genes involved in de novo lipogenesis and chylomicron formation (1, 17). Thus, we focused on documenting gene expression complementing dynamic metabolic data and hypothesized that fructose regulates in vivo the small intestinal expression of representative genes involved in lipogenesis as well as in apolipoprotein synthesis and also that regulation is dependent on GLUT5-mediated absorption and on KHK-mediated metabolism. We also noted that the identity of the cell type(s) capable of fructose-induced lipogenesis is unknown. Hence, we hypothesized that fructose regulation of lipogenic genes occurs specifically in absorptive ENT and not in secretory and stem cells. We tested this hypothesis by exposing intestinal organoids enriched in specific cell types to fructose at concentrations previously shown to upregulate fructolytic genes in vitro (12, 21, 22).

Materials and Methods

Animals

All procedures conducted in this study were approved by the Institutional Animal Care and Use Committee, New Jersey Medical School, Rutgers University. Tissues were harvested from adult, wild-type (WT; C57BL/6; Taconic Laboratories), Slc2a5-KO, or Khk-KO [both Khk isoforms A and B were ablated globally (1)] male mice. Mice were maintained under controlled temperatures (24°C) with a 12-h light–dark cycle and had free access to water and food (LabDiet Rodent 5001; Purina Mills).

Experimental design

Expression of lipogenic genes in different tissues

Samples from the liver, lung, skeletal muscle, small intestine (mucosal scrapes), and kidney were harvested from 8-wk-old WT male mice [n = 5, except kidney (n = 4)] and then immediately frozen and stored at −80°C for future analyses of gene expression.

Effect of fructose metabolism and absorption on in vivo expression of lipogenic genes in the gut

The experimental design and groups used in the in vivo study were previously described (23). Briefly, WT, Slc2a5-KO, and Khk-KO male, 5-wk-old mice were each randomly divided into 2 groups with access to food and water. They were then lightly anesthetized prior to gavage with 15% fructose (2 mL/100 g body weight, which is ∼0.3 mL per mouse) or glucose (control) solutions; a “naïve” control was not evaluated because expression of all fructolytic enzymes required for synthesis of precursors of fructose-derived lipids was similar between glucose- and lysine-perfused intestines (1). Gavage feeding occurred twice a day for 2.5 d (n = 4–5), and mice were killed after a single gavage on the third day. The dose is roughly similar to humans drinking 3–4 cans of regular soda each day for 2.5 d. After gavage feeding, mice were returned to their cages each time, and they continued to have access to water and food (LabDiet Rodent 5001: 23.9% crude protein, 5.0% fat, 5.1% crude fiber, 48.7% nitrogen-free extract, and 7.0% ash; Purina Mills). During the 2.5 d, mice exhibited normal growth and body weight for their age. Initial body weights, final body weights, and feeding rates were not affected by diet or genotype (initial body weight for all mice: 20.3 ± 0.1 g; final body weight: 21.0 ± 0.3 g; feeding rate: 4.8 ± 0.1 g mouse−1 d−1). After the mice were killed, 6–8 cm of the proximal small intestine was harvested, the mucosa was scraped ice-cold, and scrapes were stored at −80°C.

Effect of fructose on expression of lipogenic genes in organoids enriched in different cell types

Generation of intestinal organoids

Intestinal crypts were isolated from the proximal intestine of WT, Slc2a5-KO, and Khk-KO male, 5- to 8-wk-old mice fed a commercial, nonpurified rodent diet. Crypts were then cultured as previously described (12, 21) using a crypt culture medium (CCM) described in Supplemental Table 1.

Directed differentiation of intestinal stem cells

After 2–3 d in CCM, intestinal crypts grew into typical (TYP) organoids. Organoids cultured throughout in CCM (without Wnt or Notch inhibitors or activators) are referred to as TYP organoids because these have all cell types represented in roughly the same proportion as that found in vivo. On the third day, organoids were directed either to remain mostly undifferentiated and become enriched in stem cells or to differentiate and become enriched mostly in ENT, GOB, or PAN cells. Directed differentiation was achieved by activating and/or inhibiting Wnt and Notch signaling pathways (12, 21) (Supplemental Table 2). Our previous studies showed that directed differentiation resulted in organoids composed mostly of ENT (∼90% enrichment), GOB (80%), PAN (65%), or ISC (90%) cells.

Identifying the intestinal cell type involved in lipogenesis

After 2–3 d in differentiation media when enrichment in specific cell types was confirmed by biomarkers, organoids were treated with 10 mM fructose or glucose (control) for 24 h (12). We previously showed that rats fed high-fructose (65%) pellets had luminal concentrations of 26 mM (24); hence, 10 mM is physiological when mice are fed fructose at lower concentrations. Organoids were then harvested and stored at −80°C for subsequent analysis of mRNA expression.

Real-Time qPCR

Total RNA was extracted from tissues or organoids using TRIzol (Invitrogen) and then purified with an RNeasy micro kit (Qiagen) as previously described (12). After analyses of several typical housekeeping genes (Supplemental Figure 1), 18S rRNA was selected to normalize expression among liver, lung, intestine, skeletal muscle, and kidney, and β-actin was used for intestinal cell types. Primer sequences (Integrated DNA Technologies) are listed in Supplemental Table 3.

Western blot analysis

Briefly, after perfusion, the intestinal mucosa was immediately scraped from 10 cm of jejunum, snap-frozen in liquid nitrogen, and then stored at −80°C [for details, see Cui et al. (25)]. The mucosa was subsequently placed in 20 mL of ice-cold mannitol/Tris buffer (300 mM mannitol, 1 mM Tris·HCl, pH 7.4) and homogenized with a Polytron homogenizer (setting 9) for 30 s. Total protein [40 μg (Bio-Rad Laboratories)] from homogenate samples was mixed with 2 × Laemmli sample buffer (Sigma-Aldrich) and run on a 10% SDS-PAGE using a Mini-PROTEAN II cell (Bio-Rad Laboratories) for 2 h. The proteins were transferred onto a nitrocellulose membrane (Amersham Biosciences) for 1 h at room temperature using the model 6000 Electroblotter (E&K Scientific Products). Membranes were probed with primary antibody against SCD1 (1:1000; Sigma Aldrich), and all membranes were subsequently stripped and reprobed with primary antibody against the housekeeping protein β-actin (1:1000; Millipore).

Statistical analyses

Data are presented as means ± SEs. Differences of expression among different tissues and among different organoid types were analyzed with 1-way ANOVA. A multiple post hoc test (Tukey's) was done to determine significant differences between means. The effect of fructose on expression for each tissue or organoid was also analyzed by 1-way ANOVA. Differences were considered significant at P ≤ 0.05 (GraphPad Prism 8; GraphPad Software).

Results

Expression of lipogenic genes in the gut relative to those in other organ systems

Relative expression levels of Srebf1, normalized to those in the small intestine, were generally similar among the various tissues except for skeletal muscle, which expressed approximately half of those in other organs (Figure 1). Acaca was also relatively similarly expressed among different organ systems, with renal, lung, and liver expression being 3- to 5-fold greater relative to the intestine and muscle. Both Fasn and Scd1 were expressed in the liver 2- to 500-fold greater than in all other tissues. Although modest relative to that in the liver, absolute expression in the gut of Fasn and Scd1 was still plentiful because an estimate of RNA quantity, the quantification cycle (Cq), was ∼24 for both genes, whereas housekeeping gene Cqs were ∼12–18 (Supplemental Figure 1), and those of very-low-abundance genes were ∼32–35. Expression of Dgat1 and Apoa4 was at least 10-fold greater in the small intestine compared with all other tissues. Apob expression in the liver and small intestine was 4- to 20-fold greater than that in lung, skeletal muscle, and kidney. Expression levels of Gk were >5-fold in the kidney compared with other tissues, although modest amounts were expressed in the small intestine. As expected for gluconeogenic tissues, G6pc expression was >20-fold in the liver and kidney compared with other tissues (Figure 1). Mlxipl expression was 3- to 30-fold greater in the liver compared with other tissues. Thus, in the small intestine, genes involved in fatty acid synthesis were expressed modestly, whereas those involved in triglyceride synthesis and chylomicron packaging were expressed abundantly. For comparison, expression of genes (Slc5a1 and Slc2a5) coding for sugar transporters was generally abundant in the small intestine compared with all other tissues (Supplemental Figure 2A1, A2).

FIGURE 1.

FIGURE 1

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Expression of representative lipogenic genes in different organ systems. Tissues from the small intestine, liver, lung, skeletal muscle, and kidney were obtained from WT mice fed a nonpurified diet. mRNA concentrations of genes involved in lipogenesis and apolipoprotein synthesis as well as of G6pc, Slc5a1, Mlxipl, and Slc2a5 mRNA were normalized to those in the small intestine ( = 1.0). Results are means ± SEMs, n = 4–5. Different superscripts indicate significantly different means, where superscripts a > b > c. P ≤ 0.05, as determined by 1-way ANOVA. Rn18s was used as housekeeping reference gene (Supplemental Figure 1). Acaca, acetyl-CoA carboxylase 1; Apoa4, apolipoprotein A-4; Apob, apolipoprotein B; Dgat1, diacylglycerol acyltransferases; Fasn, fatty acid synthase; Gk, glycerol kinase; G6pc, glucose-6-phosphatase catalytic subunit; Mlxipl, MLX-interacting protein-like (also known as carbohydrate response element-binding protein); Scd1, stearoyl-CoA desaturase 1; Srebf1, sterol regulatory element-binding transcription factor 1; WT, wild type.

Fructose-induced increases in lipogenic gene expression depend on fructose transport and metabolism

In WT mice, fructose increased by 3- to 20-fold the small intestinal expression of all representative genes (Srebf1, Acaca, Fasn, Scd1, and Dgat1) involved in lipogenesis and triglyceride synthesis (Figure 2A–E). Deletion of Slc2a5 and of Khk each abrogated the fructose effect in all these genes. Compared with WT mice, deletion of Khk reduced by >10-fold the expression of Acaca and Fasn. The pattern of expression of Apoa4, Apob, and Gk (Figure 2F and G) was similar in that fructose upregulated by 3- to 15-fold their mRNA concentrations in WT, but there was no fructose upregulation in Slc2a5-KO and Khk-KO mice. In fact, Gk expression increased almost 3-fold with glucose in Slc2a5-KO mice compared with glucose in WT.

FIGURE 2.

FIGURE 2

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Fructose increased expression of lipogenic genes in gavage-fed WT mice. Jejunal mucosa were obtained from WT, Slc2a5-KO, and Khk-KO mice gavaged with Fru or Glu (control) solutions twice a day for 2.5 d. Expression was normalized to those of Glu-fed WT mice ( = 1.0). Results are means ± SEMs, n = 4–5. *Different from Glu, P < 0.05. Fru, fructose; Glu, glucose; KO, knockout; WT, wild type.

As in previous work (1), fructose increased by 3.5-fold G6pc expression in WT but not in Slc2a5-KO and Khk-KO mice (Figure 2I). Expression of Mlxipl, which regulates glycolysis, did not increase with fructose feeding and was 2- to 5-fold greater in the glucose-fed Slc2a5-KO and Khk-KO mice (Figure 2J). Expression of the nonlipogenic, nonfructolytic gene Slc5a1 was not affected by fructose feeding, suggesting that the fructose effect on lipogenic and apolipoprotein genes is specific (Supplemental Figure 2B1, B2). Slc2a5 expression increased with fructose in WT but not in Khk-KO mice, as previously shown (23).

Compared with those gavaged with glucose, protein concentrations of SCD1 increased in WT mice gavaged with fructose (Figure 3), a finding similar to that found for mRNA concentrations. On the other hand, deletion of Slc2a5 (Figure 3A) and Khk (Figure 3B) abolished the effects of fructose on SCD1, suggesting that the fructose-induced increase in expression required both GLUT5-mediated fructose transport and KHK-mediated metabolism.

FIGURE 3.

FIGURE 3

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Fructose increased SCD1 protein concentrations in gavage-fed WT but not in Slc2a5-KO and Khk-KO mice. (A) Protein expression levels by Western blot of SCD1 in intestinal mucosa from WT and Slc2a5-KO mice gavage fed with fructose or glucose (control) solutions twice a day for 2.5 d. (B) Protein expression levels of SCD1 in WT and Khk-KO mice. A duplicate Western blot depicted similar results. F, fructose; G, glucose; KO, knockout; WT, wild type.

Directed differentiation of organoids enriched in specific cell types

The mRNA concentrations of distinct biomarkers confirmed the differentiation of stem cells into specific cell types and were compared with those of TYP (control, contains all cell types) organoids (Figure 4A–D). Expression of leucine-rich repeat-containing G protein-coupled receptor-5 (Lgr5, a stem cell biomarker) was 4- to 50-fold greater in organoids directed to consist mostly of stem cells. Sucrase isomaltase (Si, a biomarker of ENTs) expression confirmed the enrichment of ENTs in organoids directed to differentiate primarily to this cell type. Si expression was also high in TYP organoids (∼0.3-fold compared with ENT), which also contained many ENTs. Expression of mucin 2 (Muc2, biomarker of GOB) was almost 300-fold greater in organoids directed to differentiate into GOB compared with TYP and to other organoids. Expression of the Paneth cell biomarker lysozyme (Lyz) confirmed the enrichment of PAN in organoids directed to differentiate to this cell type (∼10-fold compared with that in TYP). The distinct morphology of each organoid enriched in different cell types (Figure 4E) was similar to that previously shown (22). Thus, the organoids were each enriched in a specific cell type as directed.

FIGURE 4.

FIGURE 4

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Expression levels of biomarkers of different cell types and brightfield images of organoids. Expression of biomarkers (A. Lgr5, B. Si, C. Muc2, D. Lyz) was normalized to that in TYP organoids ( = 1.0). Results are means ± SEMs, n = 3–6. Different superscripts indicate significantly different means, where superscripts a > b > c. P ≤ 0.05, as determined by 1-way ANOVA. (E) Representative brightfield images (left to right) of crypts (arrow) harvested from mucosa; TYP organoids; as well as organoids enriched in ISCs, ENTs, and PAN and GOB cells. Images are shown at 20× magnification. ENT, enterocyte; GOB, goblet; ISC, intestinal stem cell; L, lumen; PAN, Paneth; TYP, typical.

Fructose-induced expression of lipogenic genes occurs only in enterocytes in WT mice

Because fructose stimulated the mRNA expression levels of lipogenic genes in vivo in WT mice, we next investigated whether fructose stimulation of lipogenic genes was specific to one or more cell types. Typical organoids contain cell types at proportions similar to those in vivo; thus, they contain mainly ENTs and respond similarly to ENT-enriched organoids (12, 22).

With the exception of Acaca, which increased in expression with fructose incubation in 3 cell types, the fructose-induced upregulation of mRNA expression of lipogenic genes was observed only in typical and in ENT organoids (Figure 5A–E). This suggested that the effect of fructose on gene expression depended on cell type. Fructose incubation increased expression of lipogenic genes by 1.5- to 5-fold in TYP and by 2- to 4-fold in ENT organoids. It is unclear why fructose increased Acaca expression in several cell types, but this gene, which is expressed in human intestinal epithelial cells, is regulated transcriptionally by multiple promoters that modulate Acaca mRNA abundance as a general response to nutritional status (17).

FIGURE 5.

FIGURE 5

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Fructose increased expression of lipogenic genes only in enterocytes. Each organoid was treated with 10 mM Glu (control) or 10 mM Fru for 24 h. Concentrations of mRNA in all organoids were normalized to those in TYP organoids treated with 10 mM Glu ( = 1.0). Results are means ± SEMs, n = 3–6. *Different from Glu, P < 0.05; +P = 0.06. ENT, enterocyte; Fru, fructose; Glu, glucose; GOB, goblet; ISC, intestinal stem cell; PAN, Paneth; TYP, typical.

The pattern of expression of Apoa4, Apob, and Gk (Figure 5F–H) was similar to that of lipogenic genes in that fructose increased expression by 2- to 4-fold only in TYP and ENT organoids. Confirming previous work, fructose increased G6pc expression in virtually all cell types except ISC. Fructose had no effect on Mlxipl expression (Figure 5J). In summary, expression of most genes involved in lipogenesis and apolipoprotein synthesis increased with fructose incubation primarily in ENT and TYP organoids because ENT cells also comprised most of the cells that constituted TYP organoids. The fructose effect on lipogenic genes was specific for ENTs because expression of Slc2a5 increased with fructose in all differentiated cell types and also because Slc5a1 expression did not change with fructose in any cell type (Supplemental Figure 2C1, C2).

Slc2a5 deletion prevents the majority of fructose effects in enterocytes

Because fructose effects on expression of enzymes involved in lipogenesis and apolipoprotein synthesis were observed only in ENT, we evaluated the role of Slc2a5 deletion on fructose effects only in ENT and ISC (as control) organoids. The enrichment of these organoids in ENT and ISC biomarkers is shown in Supplemental Figure 3. Fructose effects on expression of most, but not all, genes involved in lipogenesis and lipoprotein synthesis were prevented by the deletion of Slc2a5 (Supplemental Figure 4). Fructose-induced increases in Srebf1, Fasn, Scd1, Gk, and Apob were abolished in ENTs of Slc2a5-KO mice. In contrast to these genes, mRNA concentrations of Acaca, Dgat1, and Apoa4 increased modestly with fructose. Fructose increased G6pc expression in Slc2a5-KO mice—a finding similar to that of our previous study (12). Mlxipl (Supplemental Figure 4) and Slc5a1 (not shown) expression was not significantly affected by fructose. Slc5a1 is expressed mainly in differentiated cells and is sometimes used as a biomarker for differentiation (12).

Khk deletion abolishes fructose effects

The enrichment of these ENT and ISC organoids in ENT and ISC, respectively, biomarkers is shown in Supplemental Figure 5. Whereas deletion of Slc2a5 abolished fructose effects in most, but not all, lipogenic genes, deletion of Khk abolished fructose effects in all genes involved in lipogenesis and lipoprotein synthesis (Supplemental Figure 6). Expression of G6pc, Mlxipl, and Slc5a1 genes not directly involved in lipogenesis was not significantly affected by fructose in these mice.

Discussion

The main findings in our study are that 1) fructose induces the expression of almost all genes involved in lipid and apolipoprotein synthesis in the small intestine, 2) these effects are entirely dependent on fructose metabolism, and 3) fructose-induced expression is limited to differentiated ENTs.

Baseline expression of lipogenic and lipoprotein genes in the small intestine

Although the liver and adipose tissues are known sites of lipogenesis, no studies have directly compared expression of genes involved in lipid and apolipoprotein synthesis among other organ systems. These genes were found expressed in the rodent and human kidney (26, 27), lungs (28, 29), small intestine (30–32), and even skeletal muscle (33), but there were no comparisons within the same study. Our findings indicate that in mice fed a commercial rodent diet, the baseline, unstimulated expression of almost all genes involved in triglyceride and apolipoprotein synthesis is modest to abundant in the small intestine compared with other organ systems.

Dietary fructose upregulates expression of lipogenic genes in vivo

Increased intestinal mRNA expression of lipogenic and lipoprotein genes likely precedes the intestinal conversion, in humans and animal models, of fructose to lipids as well as to lipid precursors (3, 5–9). It also likely precedes the marked increases in blood concentrations of triglyceride-rich, intestine-specific APOB48 following fructose intake (8, 9), although APOB48 concentrations are also affected by clearance rates in vivo (34). It is possible that the already substantial fructose-induced increase in mRNA concentrations of lipogenic genes after 2.5 d may not have reached a final, stable point and could have increased further. Fructose also increases hepatic expression of genes involved in lipogenesis (4, 16, 35–39). In the liver, increased expression induced by fructose of genes involved in lipid and apolipoprotein synthesis has been shown to precede increased hepatic lipid production (16, 38–40). Fructose stimulates the expression of these lipogenic genes in rat liver via the transcription factor(s) sterol regulatory element-binding transcription factor 1c (SREBP-1c) and/or carbohydrate response element-binding protein (ChREBP) encoded by Mlxipl (16, 36, 40) and also by other transcription factors (41, 42). Hyperinsulinemia arising from chronic consumption of high-fructose diets may confound our findings on increases in intestinal Srebf1 expression that is regulated by insulin (17). Acute fructose intakes in vivo, however, either have no effect on or decrease postprandial insulin concentrations (43); thus, our findings may not be compromised by perturbations in insulin concentrations.

GLUT5-mediated transport from the lumen and KHK-mediated metabolism have been shown to be essential in fructose-induced changes in fructolytic enzyme expression in the liver, small intestine, and kidney in vivo (1, 23, 35, 37). For dietary fructose to regulate gene expression in vivo, ingested fructose must be absorbed by its transporter GLUT5 and not merely “tasted” by intestinal sweet receptors that sense the nutrient composition of luminal contents (44). The fructose-induced increases in lipogenic and apolipoprotein gene expression require fructose to be metabolized to fructose-1-phosphate. Thus, deletion of Khk can prevent fructose-induced increases in intestinal lipogenesis. Likewise, Khk-KO mice are protected from fructose-induced hepatic steatosis and insulin resistance (39, 45).

The cell type mediating fructose-induced upregulation of lipogenic genes

Although dietary fructose upregulates the in vivo expression of lipogenic and apolipoprotein genes, an unresolved question in our understanding of gut adaptation is whether this fructose-induced upregulation is characteristic of all, or only of a subset of, epithelial cell types lining the intestinal crypts and villi. Because absorptive ENTs make up ∼75–80% of small intestinal cells in vivo, ENTs are the likely drivers of fructose-regulated lipogenic genes; however, it remains unclear 1) whether other cell types that make up the intestinal epithelium express lipogenic genes and 2) whether the lipogenic genes in those cells are responsive to fructose. A similar question has been asked regarding sugar-processing genes such as Si, Slc2a5, and Khk that in vivo are upregulated by dietary fructose, resulting in increased absorption and portal blood concentrations of fructose (1, 46). Using a similar specialized organoid culture method, we then demonstrated that in all cell types, except stem cells, fructose upregulates expression of these genes. Although differentiation from stem cells is adequate for fructose-induced upregulation of fructolytic genes (12), further specialized differentiation into absorptive ENTs seems to be required for fructose-induced upregulation of genes involved in lipid and apolipoprotein synthesis. Because organoids from nondiabetic mice were incubated with fructose and were not exposed to changes in insulin concentrations, fructose effects on intestinal Srebf1 and other lipogenic genes may be independent of insulin. The liver is also highly responsive to dietary fructose concentrations, but it is not clear which of the 4 major hepatic cell types (hepatocytes, hepatic stellate cells, Kupffer cells, or liver sinusoidal endothelial cells) is or are involved in adapting to fructose.

Although Slc2a5 deletion resulted in a global in vivo inhibition of fructose-induced upregulation of lipogenic genes in gavage-fed mice, it did not inhibit fructose-induced in vitro upregulation of lipogenic genes Acaca, Dgat1, and Apoa4 in organoids. This is because Slc2a5 is apical and mediates the transport of dietary fructose from the lumen into the cell, so without Slc2a5, fructose cannot be absorbed in vivo. Because organoids develop with the lumen inside, organoid incubation with fructose exposes only the basolateral side, where the basolateral fructose transporter GLUT2 can mediate the transport of fructose from the medium into the cell. It is possible that fructose in the organoid media absorbed by basolateral GLUT2 was sufficient to upregulate Acaca, Dgat1, and Apoa4 in ENT organoids. Expression in organoids of Srebf1, Fasn, Scd1, Gk, and Apob did not increase with fructose because these genes may require Slc2a5 not as a transporter but, rather, as a transceptor sensing intracellular fructose and affecting intracellular signals regulating expression of these genes (13). In our previous organoid work, several fructolytic genes were also thought to depend on Slc2a5 as a transceptor and not as a transporter (12).

These directed organoids contain other cell types that could potentially confound our findings. However, each organoid type is more highly enriched in a specific cell compared to in situ populations in the small intestine, providing us an experimental approach able to estimate mRNA expression in different cell types—an objective that would be difficult to pursue in vivo and even in vitro because isolated intestinal cells have extremely short life spans (∼1 h) unless cultured in CCM and Matrigel into organoids (22).

Conclusions

Our study extends previous findings of small intestinal de novo lipogenesis from dietary fructose. Enhanced expression of genes involved in this process that takes place only in enterocytes requires fructose metabolism, suggesting that fructose metabolites are likely precursors of lipids incorporated in chylomicrons after fructose feeding (5–7).

Supplementary Material

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Acknowledgments

We are deeply indebted to Veronique Douard (National Institute of Agronomic Research, France) for reviewing earlier versions of the manuscript and to Danielle Harlan, Yaoxi Li, Ian Nadler, and John Veltri for laboratory help. The authors’ responsibilities were as follows—RPF: conceptualized the project; AA-J, CRP, SCP, KK, SY, NG, and RPF: designed the experiments; AA-J, CRP, RJS, ER, MA, JG, SCP, KK, SY, and NG: conducted the experiments; AA-J, JG, and RJS: maintained the animals; AA-J, RJS, ER, MA, and JG: analyzed the data; AA-J, wrote the manuscript; AA-J, CRP, SCP, KK, SY, NG, and RPF: revised the manuscript; RPF, finalized the manuscript; and all authors: read and approved the final manuscript.

Notes

This work was supported by National Science Foundation grants IOS-1456673 and 1754783 (RPF) and by NIH grants R01-DK-102934 (NG) and R01-AT010243 (NG and RPF).

Author disclosures: The authors report no conflicts of interest.

Supplemental Tables 1–3 and Supplemental Figures 1–6 are available from the “Online Supporting Material” link in the online posting of the article and from the same link in the online table of contents at http://academic.oup.com/jn/.

Abbreviations used: CCM, crypt culture medium; Cq, quantification cycle; DGAT1, diacylglycerol acyltransferase 1; ENT, enterocyte; Gk, glycerol kinase; GLUT, glucose transporter; GOB, goblet cell; G6pc, gene encoding for glucose-6-phosphatase catalytic subunit; ISC, intestinal stem cell; KHK, ketohexokinase; KO, knockout; PAN, Paneth cell; SCD1, stearoyl-CoA desaturase 1; Si, gene encoding for sucrase isomaltase; Slc, solute carrier family; TYP, typical; WT, wild type.

Contributor Information

Arwa Al-Jawadi, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Chirag R Patel, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Reilly J Shiarella, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Emmanuellie Romelus, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Madelyn Auvinen, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Joshua Guardia, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Sarah C Pearce, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Kunihiro Kishida, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

Shiyan Yu, Department of Biological Sciences, Life Science Center, Rutgers University, Newark, NJ, USA.

Nan Gao, Department of Biological Sciences, Life Science Center, Rutgers University, Newark, NJ, USA.

Ronaldo P Ferraris, Department of Pharmacology, Physiology & Neuroscience, New Jersey Medical School, Rutgers University, Newark, NJ, USA.

References

  • 1. Patel C, Douard V, Yu S, Tharabenjasin P, Gao N, Ferraris RP. Fructose-induced increases in expression of intestinal fructolytic and gluconeogenic genes are regulated by GLUT5 and KHK. Am J Physiol Regul Integr Comp Physiol. 2015;309:R499–509. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 2. Jensen T, Abdelmalek MF, Sullivan S, Nadeau KJ, Green M, Roncal C, Nakagawa T, Kuwabara M, Sato Y, Kang DHet al.. Fructose and sugar: a major mediator of non-alcoholic fatty liver disease. J Hepatol. 2018;68:1063–75. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 3. Jang C, Hui S, Lu W, Cowan AJ, Morscher RJ, Lee G, Liu W, Tesz GJ, Birnbaum MJ, Rabinowitz JD. The small intestine converts dietary fructose into glucose and organic acids. Cell Metab. 2018;27:351–61 e3. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 4. Steenson S, Umpleby AM, Lovegrove JA, Jackson KG, Fielding BA. Role of the enterocyte in fructose-induced hypertriglyceridaemia. Nutrients. 2017;9:E349. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5. Egli L, Lecoultre V, Theytaz F, Campos V, Hodson L, Schneiter P, Mittendorfer B, Patterson BW, Fielding BA, Gerber PAet al.. Exercise prevents fructose-induced hypertriglyceridemia in healthy young subjects. Diabetes. 2013;62:2259–65. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. Theytaz F, de Giorgi S, Hodson L, Stefanoni N, Rey V, Schneiter P, Giusti V, Tappy L. Metabolic fate of fructose ingested with and without glucose in a mixed meal. Nutrients. 2014;6:2632–49. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 7. Surowska A, De Giorgi S, Theytaz F, Campos V, Hodson L, Stefanoni N, Rey V, Schneiter P, Laville M, Schneiter Pet al.. Effects of roux-en-Y gastric bypass surgery on postprandial fructose metabolism. Obesity (Silver Spring). 2016;24:589–96. [DOI] [PubMed] [Google Scholar]
  • 8. Xiao C, Dash S, Morgantini C, Lewis GF. Novel role of enteral monosaccharides in intestinal lipoprotein production in healthy humans. Arterioscler Thromb Vasc Biol. 2013;33:1056–62. [DOI] [PubMed] [Google Scholar]
  • 9. Haidari M, Leung N, Mahbub F, Uffelman KD, Kohen-Avramoglu R, Lewis GF, Adeli K. Fasting and postprandial overproduction of intestinally derived lipoproteins in an animal model of insulin resistance: evidence that chronic fructose feeding in the hamster is accompanied by enhanced intestinal de novo lipogenesis and ApoB48-containing lipoprotein overproduction. J Biol Chem. 2002;277:31646–55. [DOI] [PubMed] [Google Scholar]
  • 10. Goncalves MD, Lu C, Tutnauer J, Hartman TE, Hwang SK, Murphy CJ, Pauli C, Morris R, Taylor S, Bosch Ket al.. High-fructose corn syrup enhances intestinal tumor growth in mice. Science. 2019;363:1345–9. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 11. Schwitalla S, Fingerle AA, Cammareri P, Nebelsiek T, Goktuna SI, Ziegler PK, Canli O, Heijmans J, Huels DJ, Moreaux Get al.. Intestinal tumorigenesis initiated by dedifferentiation and acquisition of stem-cell-like properties. Cell. 2013;152:25–38. [DOI] [PubMed] [Google Scholar]
  • 12. Kishida K, Pearce SC, Yu S, Gao N, Ferraris RP. Nutrient sensing by absorptive and secretory progenies of small intestinal stem cells. Am J Physiol Gastrointest Liver Physiol. 2017;312:G592–605. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 13. Ferraris RP, Choe JY, Patel CR. Intestinal absorption of fructose. Annu Rev Nutr. 2018;38:41–67. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 14. Douard V, Ferraris RP. The role of fructose transporters in diseases linked to excessive fructose intake. J Physiol. 2013;591:401–14. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 15. Dekker MJ, Su Q, Baker C, Rutledge AC, Adeli K. Fructose: a highly lipogenic nutrient implicated in insulin resistance, hepatic steatosis, and the metabolic syndrome. Am J Physiol Endocrinol Metab. 2010;299:E685–94. [DOI] [PubMed] [Google Scholar]
  • 16. Miyazaki M, Dobrzyn A, Man WC, Chu K, Sampath H, Kim HJ, Ntambi JM. Stearoyl-CoA desaturase 1 gene expression is necessary for fructose-mediated induction of lipogenic gene expression by sterol regulatory element-binding protein-1c-dependent and -independent mechanisms. J Biol Chem. 2004;279:25164–71. [DOI] [PubMed] [Google Scholar]
  • 17. Gutierrez-Repiso C, Rodriguez-Pacheco F, Garcia-Arnes J, Valdes S, Gonzalo M, Soriguer F, Moreno-Ruiz FJ, Rodriguez-Canete A, Gallego-Perales JL, Alcain-Martinez Get al.. The expression of genes involved in jejunal lipogenesis and lipoprotein synthesis is altered in morbidly obese subjects with insulin resistance. Lab Invest. 2015;95:1409–17. [DOI] [PubMed] [Google Scholar]
  • 18. Gaby AR. Adverse effects of dietary fructose. Altern Med Rev. 2005;10:294–306. [PubMed] [Google Scholar]
  • 19. Hirahatake KM, Meissen JK, Fiehn O, Adams SH. Comparative effects of fructose and glucose on lipogenic gene expression and intermediary metabolism in HepG2 liver cells. PLoS One. 2011;6:e26583. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 20. Cheng D, Iqbal J, Devenny J, Chu CH, Chen L, Dong J, Seethala R, Keim WJ, Azzara AV, Lawrence RMet al.. Acylation of acylglycerols by acyl coenzyme A:diacylglycerol acyltransferase 1 (DGAT1): functional importance of DGAT1 in the intestinal fat absorption. J Biol Chem. 2008;283:29802–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 21. Yin X, Farin HF, van Es JH, Clevers H, Langer R, Karp JM. Niche-independent high-purity cultures of Lgr5+ intestinal stem cells and their progeny. Nat Methods. 2014;11:106–12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 22. Pearce SC, Al-Jawadi A, Kishida K, Yu S, Hu M, Fritzky LF, Edelblum KL, Gao N, Ferraris RP. Marked differences in tight junction composition and macromolecular permeability among different intestinal cell types. BMC Biol. 2018;16:19. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 23. Patel C, Douard V, Yu S, Gao N, Ferraris RP. Transport, metabolism, and endosomal trafficking-dependent regulation of intestinal fructose absorption. FASEB J. 2015;29:4046–58. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 24. Jiang L, Ferraris RP. Developmental reprogramming of rat GLUT-5 requires de novo mRNA and protein synthesis. Am J Physiol Gastrointest Liver Physiol. 2001;280:G113–20. [DOI] [PubMed] [Google Scholar]
  • 25. Cui XL, Schlesier AM, Fisher EL, Cerqueira C, Ferraris RP. Fructose-induced increases in neonatal rat intestinal fructose transport involve the PI3-kinase/Akt signaling pathway. Am J Physiol Gastrointest Liver Physiol. 2005;288:G1310–20. [DOI] [PubMed] [Google Scholar]
  • 26. Wang Z, Jiang T, Li J, Proctor G, McManaman JL, Lucia S, Chua S, Levi M. Regulation of renal lipid metabolism, lipid accumulation, and glomerulosclerosis in FVBdb/db mice with type 2 diabetes. Diabetes. 2005;54:2328–35. [DOI] [PubMed] [Google Scholar]
  • 27. Herman-Edelstein M, Scherzer P, Tobar A, Levi M, Gafter U. Altered renal lipid metabolism and renal lipid accumulation in human diabetic nephropathy. J Lipid Res. 2014;55:561–72. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 28. Zhang F, Pan T, Nielsen LD, Mason RJ. Lipogenesis in fetal rat lung: importance of C/EBPα, SREBP-1c, and stearoyl-CoA desaturase. Am J Respir Cell Mol Biol. 2004;30:174–83. [DOI] [PubMed] [Google Scholar]
  • 29. Antony N, Weir JR, McDougall AR, Mantamadiotis T, Meikle PJ, Cole TJ, Bird AD. cAMP response element binding protein 1 is essential for activation of steroyl co-enzyme a desaturase 1 (Scd1) in mouse lung type II epithelial cells. PLoS One. 2013;8:e59763. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 30. Al-Dwairi A, Brown AR, Pabona JM, Van TH, Hamdan H, Mercado CP, Quick CM, Wight PA, Simmen RC, Simmen FA. Enhanced gastrointestinal expression of cytosolic malic enzyme (ME1) induces intestinal and liver lipogenic gene expression and intestinal cell proliferation in mice. PLoS One. 2014;9:e113058. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 31. Levy E. Insights from human congenital disorders of intestinal lipid metabolism. J Lipid Res. 2015;56:945–62. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 32. Yen CL, Nelson DW, Yen MI. Intestinal triacylglycerol synthesis in fat absorption and systemic energy metabolism. J Lipid Res. 2015;56:489–501. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 33. Funai K, Song H, Yin L, Lodhi IJ, Wei X, Yoshino J, Coleman T, Semenkovich CF. Muscle lipogenesis balances insulin sensitivity and strength through calcium signaling. J Clin Invest. 2013;123:1229–40. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 34. Tremblay AJ, Lamarche B, Hogue JC, Couture P. Effects of ezetimibe and simvastatin on apolipoprotein B metabolism in males with mixed hyperlipidemia. J Lipid Res. 2009;50:1463–71. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 35. Ishimoto T, Lanaspa MA, Le MT, Garcia GE, Diggle CP, Maclean PS, Jackman MR, Asipu A, Roncal-Jimenez CA, Kosugi Tet al.. Opposing effects of fructokinase C and A isoforms on fructose-induced metabolic syndrome in mice. Proc Natl Acad Sci U S A. 2012;109:4320–5. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 36. Koo HY, Miyashita M, Cho BH, Nakamura MT. Replacing dietary glucose with fructose increases ChREBP activity and SREBP-1 protein in rat liver nucleus. Biochem Biophys Res Commun. 2009;390:285–9. [DOI] [PubMed] [Google Scholar]
  • 37. Lanaspa MA, Ishimoto T, Li N, Cicerchi C, Orlicky DJ, Ruzycki P, Rivard C, Inaba S, Roncal-Jimenez CA, Bales ESet al.. Endogenous fructose production and metabolism in the liver contributes to the development of metabolic syndrome. Nat Commun. 2013;4:2434. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 38. Softic S, Cohen DE, Kahn CR. Role of dietary fructose and hepatic de novo lipogenesis in fatty liver disease. Dig Dis Sci. 2016;61:1282–93. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 39. Softic S, Gupta MK, Wang GX, Fujisaka S, O'Neill BT, Rao TN, Willoughby J, Harbison C, Fitzgerald K, Ilkayeva Oet al.. Divergent effects of glucose and fructose on hepatic lipogenesis and insulin signaling. J Clin Invest. 2017;127:4059–74. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 40. Liu L, Wang S, Yao L, Li JX, Ma P, Jiang LR, Ke DZ, Pan YQ, Wang JW. Long-term fructose consumption prolongs hepatic stearoyl-CoA desaturase 1 activity independent of upstream regulation in rats. Biochem Biophys Res Commun. 2016;479:643–8. [DOI] [PubMed] [Google Scholar]
  • 41. Samuel VT. Fructose induced lipogenesis: from sugar to fat to insulin resistance. Trends Endocrinol Metab. 2011;22:60–5. [DOI] [PubMed] [Google Scholar]
  • 42. Kelley GL, Azhar S. Reversal of high dietary fructose-induced PPARα suppression by oral administration of lipoxygenase/cyclooxygenase inhibitors. Nutr Metab (Lond). 2005;2:18. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 43. Evans RA, Frese M, Romero J, Cunningham JH, Mills KE. Fructose replacement of glucose or sucrose in food or beverages lowers postprandial glucose and insulin without raising triglycerides: a systematic review and meta-analysis. Am J Clin Nutr. 2017;106:506–18. [DOI] [PubMed] [Google Scholar]
  • 44. Shirazi-Beechey SP, Daly K, Al-Rammahi M, Moran AW, Bravo D. Role of nutrient-sensing taste 1 receptor (T1R) family members in gastrointestinal chemosensing. Br J Nutr. 2014;111(Suppl 1):S8–15. [DOI] [PubMed] [Google Scholar]
  • 45. Miller CO, Yang X, Lu K, Cao J, Herath K, Rosahl TW, Askew R, Pavlovic G, Zhou G, Li Cet al.. Ketohexokinase knockout mice, a model for essential fructosuria, exhibit altered fructose metabolism and are protected from diet-induced metabolic defects. Am J Physiol Endocrinol Metab. 2018;315:E386–93. [DOI] [PubMed] [Google Scholar]
  • 46. Patel C, Sugimoto K, Douard V, Shah A, Inui H, Yamanouchi T, Ferraris RP. Effect of dietary fructose on portal and systemic serum fructose levels in rats and in KHK–/– and GLUT5–/– mice. Am J Physiol Gastrointest Liver Physiol. 2015;309:G779–90. [DOI] [PMC free article] [PubMed] [Google Scholar]

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