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Cellular prion protein promotes glucose uptake through the Fyn-HIF-2α-Glut1 pathway to support colorectal cancer cell survival



This article is corrected by:

  1. Errata: Correction Volume 103, Issue 3, 606, Article first published online: 4 March 2012

To whom correspondence should be addressed.
Email: gaowenchao@msn.com


Cellular prion protein (PrPc) is a glycosylphosphatidylinositol-anchored membrane protein that has various physical functions, including protection against apoptotic and oxidative stress, cellular uptake of copper ions, transmembrane signaling, and adhesion to the extracellular matrix. In this study, we show that PrPc is highly expressed in colorectal adenocarcinomas. Transcriptome profiling of PrPc-depleted DLD-1 cells revealed downregulation of glucose transporter 1 (Glut1). PrPc is shown to be involved in regulating Glut1 expression through the Fyn-HIF-2α pathway. As Glut1 is the natural transporter of glucose and is required for the high glycolytic rate seen in colorectal tumors, silencing of PrPc reduced the proliferation and survival rate of colorectal cancer cells in vitro. In vivo, knockdown of PrPc by hydrodynamic injection with a cocktail of PrPc–shRNA-encoding plasmids also inhibited tumorigenicity in a xenograft model in nude mice. In summary, our data characterize a novel molecular mechanism that links PrPc expression to the regulation of glycolysis. Targeting PrPc will therefore be a promising strategy to overcome the growth and survival advantage in colorectal tumors. (Cancer Sci 2011; 102: 400–406)

Colorectal cancer is one of the most commonly diagnosed cancers. It remains a leading cause of cancer-related death, with high risk of postoperative 5-year relapse.(1) Although conventional treatment results have improved somewhat over the past years, they are not always as effective as has traditionally been believed. The greatest advances in treating cancer have come from targeted therapies biologically engineered to attack cancer at its molecular roots and spare most of the normal tissue from damage. Several recent studies have implied that targeted therapies in conjunction with chemo- or radiotherapy are potential approaches for a rational molecular-based tumor therapy in colorectal cancer.(2,3)

The normal cellular prion protein (PrPc) is a glycoprotein highly conserved in mammalian species. Most PrPc molecules are normally localized on the cell surface, where they are attached to the lipid bilayer through a c-terminal, glycosylphosphatidylinositol anchor.(4) Prions accumulate not only in the central nervous system, but also in lymphoid organs, as has been shown in new variant and sporadic Creutzfeldt–Jakob disease patients, and in some animals.(5) A variety of functions have been proposed for PrPc, including involvement in cell death and survival, differentiation, metal ion trafficking,(6) cell adhesion,(7) and transmembrane signaling.(8)

Now, several intriguing lines of evidence have emerged indicating that PrPc may be implicated in tumor cell biology.(9–12) PrPc overexpression is correlated to the acquisition by tumor cells of a phenotype for resistance to cell death induced by tumor necrosis factor-α(13) or antitumor drugs.(14) Also, PrPc plays a fundamental role in the proliferation and metastasis of human cancer cells. PrPc may promote tumorigenesis, metastasis, proliferation, and G1/S transition in gastric cancer cells.(12) PrPc therefore has both vital and lethal functions. It seems that PrPc may serve as a promising target for novel anticancer therapies.

We show here that PrPc depletion inhibits tumor growth of colorectal cancer both in vivo and in vitro. Our data support the concept that PrPc expression is associated with malignant potential of colorectal cancer cells because it mediates the Warburg effect by regulating glucose transport expression through epigenetic activation of the Fyn–hypoxia-inducible factor (HIF)-2α pathway. Based on these data, inhibition of PrPc represents a potential avenue to inhibit growth of cancer cells which rely on increased glucose fermentation.

Materials and Methods

Cell lines.  The human colorectal carcinoma cell lines DLD-1, SW48, and CX-1, were purchased from American Type Culture Collection (Manassas, VA, USA) and were cultured under the conditions recommended by the manufacturer.

Bromodeoxyuridine (BrdU) incorporation and MTT assays.  Bromodeoxyuridine incorporation and MTT assays were carried out using a colorimetric BrdU and MTT assay (Roche, Indianapolis, IN, USA) according to the manufacturer’s instructions.

Immunoblotting.  Total protein was extracted from cells using RIPA lysis buffer (Santa Cruz Biotechnology, Santa Cruz, CA, USA). Protein extract (50 μg/lane) was electrophoresed, transferred to PVDF membranes, and incubated overnight with primary antibodies against Glut1, matrix metalloproteinase (MMP)2, MMP9, MMP11 (all Santa Cruz Biotechnology), PrPc, and HIF-2α (both Sigma Aldrich, St Louis, MO, USA). Membranes were then treated with the appropriate HRP-conjugated secondary antibodies (Invitrogen, Carlsbad, CA, USA). Detection was carried out using the reagents provided in the ECL Plus kit (GE Healthcare, Milwaukee, WI, USA).

Chromatin immunoprecipitation.  DLD-1 and SW480 cells were transfected with PrPc siRNA for 72 h prior to carrying out ChIP using antibodies against HIF-1α, HIF-2α, RNA polymerase II- or acetylated-histone H3, as described previously. A total of 1.0–2.5% of each immunoprecipitation was assayed by PCR using primers specific for a region of interest.

Glycolysis.  The glycolytic rate of the cells was determined by measuring the conversion of 5-3H-glucose to tritiated water, as described previously.(15)

Glucose uptake assay.  Cellular glucose uptake was measured by incubating cells in glucose-free RPMI-1640 with 0.2 Ci/mL [3H]2-deoxyglucose (specific activity, 40 Ci/mmol) for 60 min. After the cells were washed with ice-cold PBS, the radioactivity in the cell pellets was quantified by liquid scintillation counting.

Luciferase reporter assay.  The 2692 bp Glut1 untranslated region was cloned by PCR amplification. After digested by restriction endonucleases KpnI, HindIII, SacI, and XhoI, different fragments of the 5′-untranslated region were cloned into the pGL3-basic vector (Promega, Madison, WI, USA) and named pGL3-2692-luc, pGL3-1860-luc, and pGL3-631-luc, respectively. pGL2-4kb-luc contains 4 kb of the 5′-flanking region of the KDR gene linked to the firefly luciferase gene (generously provided by Edgar Haber, Harvard School of Public Health, Boston, MA, USA). Cells of 50% confluence in 24-well plates were transfected using Lipofectamine 2000 (Invitrogen) and the luciferase activity was measured as described previously.(16)

Immunofluorescence microscopy analysis.  Immunofluorescence microscopy analysis using antibodies against HIF-2α (Sigma) was carried out as described previously.(14)

Inhibition of PrPc and HIF-2α expression by RNA interference.  Cells (2 × 105) were seeded in 6-well plates in triplicate and after an overnight incubation, the cells were transfected with various concentrations of siRNA using HiPerfect Reagent (Qiagen, Valencia, CA, USA) as suggested by the manufacturer’s instructions. The small interference RNA used to target PRNP and HIF-2α mRNA sequences were synthesized by Qiagen.

In vitro kinase assays.  For studies of Fyn activation in the presence and absence of PrPc, in vitro kinase assays were carried out on Fyn, as previously described.(17)

Animal studies.  For in vivo treatment studies, 6-week-old NOD/SCID female mice were injected with 1.5 × 106 SW480 cells s.c. into the right flank. Approximately 12 days after inoculation, all mice developed palpable tumors. Tumor volume was calculated using the following formula: V = (DL × DS2) × π/6, where DL is the largest diameter and DS is the smallest diameter. Three human PrPc-specific shRNAs were designed as 62-mers containing a hairpin loop and cloned into pSUPER vector (Invitrogen). Injections were carried out hydrodynamically directly into the tumor mass, in a final volume of 100 mL. All experiments received prior approval from the Institutional Animal Use and Care Committee of Changzheng Hospital (Shanghai, China).

Patient study.  Patients were identified from the Changzheng Hospital records. The ethics committees of the participating hospital approved the written informed consent for the collection of tumor tissue obtained from each patient. Histological classification of colorectal carcinomas proposed by the World Health Organization was adopted here.(18) Three-micrometer sections were cut from paraffin blocks onto silanized slides, and the sections were immunostained using antibodies against PrPc (Sigma), Glut1 (BioGenex, San Ramon, CA, USA), and HIF-2α (Sigma) respectively. Expression of PrPc, Glut1, and HIF-2α was graded from 0 to 3, based on an assessment of the intensity of the reaction product, and the percentage of positive cells: score 0, negative/weak staining for all tumor cells or moderate staining <10% of tumor cells; 1, moderate staining with areas larger than 10% but <30%, or strong staining within 10% of tumor cells; 2, moderate staining with areas larger than 30% but <70%, or strong staining within 30% of tumor cells; 3, moderate staining with areas more than 70% or strong staining with areas more than 30% of all tumor cells. Pearson’s correlation coefficients were used to determine whether two factors were correlated to each other over all cases.

Statistical analysis.  Statistics were calculated by spss software (SPSS, Chicago, IL, USA). The results are presented as the mean ± SEM. Anova, Student’s t-test analysis, and Dunnett’s multiple comparison tests were used to compare mean values. A P-value of <0.05 was defined as statistical significance.


PrPc expression in colorectal carcinoma tissues.  In 23 cases of normal colorectal mucosa tissue, negative to weak membrane/cytoplasm staining of PrPc was detected (Fig. 1A). However, PrPc could be observed in 130 primary colorectal adenocarcinomas, especially in moderately differentiated (III) and undifferentiated (IV) tumors. Moreover, as shown in Figure 1, PrPc expression gradually increases according to histological grade, being highest in undifferentiated (IV) tumors (< 0.05).

Figure 1.

 Cellular prion protein (PrPc) expression levels correlate with histological grades in human colorectal carcinomas. (A) Representative immunostaining of PrPc in normal colorectal tissues and colorectal carcinomas. Scale bars = 20 μm. (B) Correlation of PrPc expression with histological grades of colorectal carcinomas (n = 130). Histological classification carried out according to World Health Organization guidelines.

Silencing of PrPc inhibits colorectal cancer cell growth in vitro.  To better understand the role of PrPc in colorectal cancer cells, we used RNA interference. As evaluated by quantitativeRT-PCR and immunoblotting, Figure 2(A,B) show the knockdown of PrPc at 48 and 72 h after transfection of DLD-1 and SW480 cells with PrPc siRNA. PrPc has been suggested to induce MMPs in tumor cells, yet silencing of PrPc did not elicit a relevant reduction in MMPs expression (Fig. 2B). However, in viability assays over a time course of 72 h, significant reduction of viability in PrPc-depleted DLD-1 and SW480 cells was observed (Fig. 2C). Furthermore, BrdU incorporation was also significantly altered up to 72 h after transfection of PrPc siRNA (Fig. 2D), suggesting that PrPc expression is essential for the survival and proliferation of colorectal cancer cells.

Figure 2.

 Cellular prion protein (PrPc) depletion suppresses colorectal cancer cell growth in vitro. After cells were transfected with PrPc siRNA over a time course of 48 and 72 h, PrPc and matrix metalloproteinase (MMP) expressions were assessed by real-time PCR (A) and immunoblotting (B) –, no treatment. The viability and proliferation rates of PrPc-depleted cells were determined by MTT (C) and bromodeoxyuridine (BrdU) incorporation assays (D). **< 0.05 versus controls.

PrPc regulates Glut1 gene expression through HIF-2α.  To find genes involved in the observed sensitization towards cell growth inhibition, we used transcriptome profiling. After 72 h of PrPc depletion in DLD-1 cells, analysis of array hybridization revealed 106 differentially expressed genes. Glut1 was further investigated because it was the most significantly changed gene in PrPc-depleted DLD-1 cells (Fig. 3A). As shown in Figure 3(B,C), Glut1 mRNA was downregulated 3.7-fold in DLD-1, and 4.2-fold in SW480 cells 72 h after PrPc silencing, according to the results obtained from immunoblot analysis. On the contrary, Glut1 expression was significantly enhanced upon pCEP4-PrPc transfection into CX-1 cells in which PrPc is negligibly expressed (Fig. S1A). In addition, relative luciferase units from pGL3-1860-luc showed higher transcriptional activity in DLD-1 cells than two other vectors, pGL3-2692-luc and pGL3-631-luc (< 0.05), whereas relative luciferase units from these three vectors showed no obvious activity in PrPc-depleted cells (Fig. 3D). Thus, Glut1 promoter drives gene expression in PrPc-expressing cells and its core promoter region may locate within the −631 and −1860 bp of the 5′-untranslated region.

Figure 3.

 Cellular prion protein (PrPc) regulates glucose transporter 1 (Glut1) expression in a hypoxia-inducible factor (HIF)-2α-dependent manner. (A) Array hybridization analysis of gene expression extracted from DLD-1 cells treated without (upper array) or with (lower array) PrPc siRNA. Glut1, HIF-1α, and HIF-2α expressions in PrPc-depleted cells were assessed by real-time PCR (B) and immunoblotting (C). (D) Relative promoter activity of pGL3-2692-luc, pGL3-1860-luc, and pGL3-631-luc in DLD-1 cells treated with or without PrPc siRNA. (E) After PrPc depletion, ChIP was carried out with HIF-1α, HIF-2α, RNA polymerase II-, acetylated-histone H3-specific antibody, or IgG as negative control. Ten percent of the chromatin input was amplified with specific primers for Glut1. −, without treatment; +, with treatment. (F) Representative immunostaining of PrPc, Glut1, and HIF-2α in colorectal carcinomas. Scale bars = 20 μm. **< 0.05 versus controls.

Glut1 promoter, which contains a typical HIF binding site, is known to respond to both hypoxia and mitochondrial inhibitors (Fig. S2). However, in ChIP-PCR assays, not HIF-1α but direct and specific binding of HIF-2α to the Glut1 locus was observed (Fig. 3E). PrPc silencing abolished the binding of HIF-2α to the Glut1 promoter (Fig. 3E) and had no impact on the expression levels of either HIF-1α or HIF-2α (Fig. 3B,C). Furthermore, we also observed decreased binding of acetylated histone H3 and RNA polymerase II to the Glut1 promoter after PrPc depletion, arguing for a condensed chromatin structure and ceased transcription (Fig. 3E).

The correlation between PrPc, Glut1, and HIF-2α expression was also evaluated in colorectal carcinoma specimens. Immunohistochemical staining for Glut1 and HIF-2α was not seen in normal colorectal mucosa (data not shown), but in 89.2% and 96.9%, respectively, of the colorectal carcinoma cells. The immunostaining of Glut1 was paralleled with that of PrPc in most colorectal carcinoma tissues. Representative immunostaining of Glut1 and PrPc are shown in Figure 3(F). Pearson’s correlation coefficient between Glut1 and PrPc was 0.910, < 0.001 (Table 1). However, differences in the expression of HIF-2α did not reach statistical significance when averaged over all samples (Fig. 3F, Table 1).

Table 1.   Correlation of cellular prion protein (PrPc) expression with glucose transporter 1 (Glut1) and hypoxia-inducible factor (HIF)-2α (n = 130)
  1. NA, not applicable.


Involvement of Fyn-HIF-2α pathway in PrPc-mediated Glut1 expression.  Recent studies have advanced the hypothesis that several signaling pathways, such as the PI3-kinase/Akt, cyclic AMP/protein kinase A, PKC, Fyn, and Erk1/2 pathways, are modulated by PrPc expression. Among them, only the Fyn pathway is constitutively activated in DLD-1 and SW480 cells, which could not be observed after PrPc siRNA treatment (Fig. 4A). In contrast, Fyn activity was greatly elevated in CX-1 cells upon ectopic expression of PrPc (Fig. S1B). When the Fyn pathway was blocked by a specific inhibitor, PP2, the binding of HIF-2α to Glut1 promoter was abolished (Fig. 4B), accompanied by downregulation in Glut1 expression (Fig. 4C). In accordance, either enhanced HIF-2α binding activity or Glut1 expression in PrPc-overexpressed CX-1 cells was prevented by treatment with PP2 and/or HIF-2α siRNA (Fig. S1C,D).

Figure 4.

 Fyn–hypoxia-inducible factor (HIF)-2α signaling pathway participates in cellular prion protein (PrPc)-mediated glucose transporter 1 (Glut1) expression. (A) PrPc-depleted DLD-1 and SW480 colorectal cancer cells were lysed and immunoprecipitated with anti-Fyn Ab, after which the immunoprecipitates were used in the autophosphorylation and substrate (enolase) phosphorylation assays. After cells were pretreated with PP2, ChIP-PCR for specific enrichment of HIF-2α (B) and immunoblotting for Glut1 (C) were carried out. (D) Cells were transfected with KDR promoter and PP2, then luciferase assays were carried out. (E) Immunocytochemistry for HIF-2α after DLD-1 cells were treated with PrPc siRNA and/or PP2. Scale bars = 20 μm. **< 0.05 versus controls.

To further investigate whether the Fyn pathway is responsible for HIF-2α activation, we measured KDR expression, a specific target gene of HIF-2α. PP2 largely inhibited both KDR promoter activity and KDR expression in DLD-1 and SW480 cells (Fig. 4D). We then attempted to evaluate the functional activity of HIF-2α upon Fyn inactivation. As shown in Figure 4(E), most HIF-2α protein was initially accumulated in the nucleus. When Fyn activity was suppressed by PrPc siRNA and/or PP2, we visualized nucleo-cytoplasmic translocation of HIF-2α in DLD-1 cells.

PrPc–HIF-2α-Glut1 axis essential for Warburg effect in colorectal cancer cells.  To test whether colorectal cancer cells show the Warburg effect, we assessed glucose consumption in DLD-1 cells. During lactic acid fermentation, 1 mol glucose is converted to 2 mol lactate and, indeed, lactate was detected to be generated at approximately double the rate of glucose consumption (Fig. 5A). The Warburg effect, defined by an increase in aerobic glycolysis, is essential for tumor cell survival and proliferation. Treatment of the glycolysis inhibitors, NaF and 2DG, reduced the viability of DLD-1 and SW480 cells in vitro in a dose-dependent fashion, as assessed by MTT assays (Fig. 5B). However, glycolysis inhibition had no impact on expression levels of either Glut1 or PrPc (Fig. 5C).

Figure 5.

 Cellular prion protein–hypoxia-inducible factor 2α–glucose transporter 1 (PrPc-HIF-2α-Glut1) axis plays a critical role in the Warburg effect of colorectal cancer cells. (A) Concentrations of lactic acid and glucose were assessed in the supernatant of DLD-1 cells at the indicated time points. After DLD-1 and SW480 cells were pretreated with (+) or without (−) NaF and 2DG, cell viability and glucose transporter 1 (Glut1) and PrPc expressions were evaluated by MTT assays (B) and immunoblotting (C). (D) The effects of PrPc siRNA, HIF-2 siRNA, and anti-Glut1 antibody on the glycolytic rate and glucose uptake in DLD-1 and SW480 cells. **< 0.05 versus controls.

To evaluate the association between Glut1 overexpression and the Warburg effect in colorectal cancer cells, we examined the effect of PrPc silencing on the glycolysis rate in DLD-1 and SW480 cells. As compared with controls, PrPc depletion significantly inhibited glycolysis as well as glucose uptake. Also, both anti-Glut1 antibody and HIF-2α siRNA exerted similar effects to PrPc siRNA in terms of anaerobic glycolysis and glucose utilization (Fig. 5D).

PrPc silencing inhibits tumorigenicity of SW480 cells in in vivo models.  To evaluate the effects of PrPc targeted therapy on colorectal tumor growth in vivo, we used PrPc shRNA cocktail in the treatment of a human colorectal xenograft model. Treatment was initiated when the tumor volume was at least 30 mm3 (typically approximately 12 days post-inoculation). We injected the plasmid cocktail hydrodynamically, using 5 μg each shRNA construct (total 15 μg) or 15 μg pSUPER control in PBS, by intratumoral injection at days 1, 3, 5, 9, 11, and 15 for a total experimental period of 15 days. Tumors from mice injected with the shRNA cocktail grew significantly slower than control injected mice. Indeed, after 35 days of treatment with shRNA, the mean tumor volume was reduced by 69.2% (P < 0.05) (Fig. 6A), and the mean tumor weight was decreased by 78.3% (P < 0.05) (Fig. 6B), suggesting that the PrPc shRNA cocktail has significant antitumor activity in colorectal cancer. Animal survival was represented by a Kaplan–Meier curve. At the end of the experiment, there was 25% survival in control animals compared to 100% in shRNA-treated animals (Fig. 6C). The difference in survival between groups was significant as indicated by the log–rank test (P = 0.0042 0.05).

Figure 6.

 Cellular prion protein (PrPc) shRNA cocktail inhibits tumor growth in vivo. NOD/SCID mice s.c. injected with SW480 cells were randomly assigned to treatment groups and injected with shRNA cocktail or control. Tumor volume (A) and weight (B) were measured at 35 days after the inoculation of shRNA cocktail or control. (C) Survival analysis is represented by a Kaplan–Meier curve. **< 0.05 versus controls. N.S, normal saline.


Cancer cells generally use glucose for aerobic glycolysis to support their rapid proliferation and expansion across the body. Aggressive tumors frequently display high rates of glycolysis even in the presence of oxygen, known as the Warburg effect, and reveal increasing dependency on this glycolytic pathway for ATP generation.(19-21) Hypoxia also has the potential to promote malignant phenotypes of tumor cells, which acquire the ability to withstand harsher microenvironments by switching to glycolysis.(22) This has led to the development of a therapeutic concept to inhibit glycolysis as a strategy to preferentially kill cancer cells. Recently, it was reported that hypoxia conditions, which positively correlate with histological grade, induce overexpression of PrPc in gastric cancer cells(23) and neuroblastoma cells.(24) We also observed that PrPc expression was closely associated with histological grading in colorectal carcinomas. Furthermore, PrPc depletion directly inhibits glucose utilization of colorectal cancer cells and thereby suppresses tumor growth both in vitro and in vivo, suggesting the possible role of PrPc in the tumor protective mechanism of glycosis. However, the exact function of PrPc still remains unclear. Several lines of evidence suggest that PrPc might have a role in the regulation of the glycolytic pathway. Lactate dehydrogenase A, a glycolytic enzyme that plays a significant role in the regulation of the glycolytic pathway, is found to be upregulated following PrPc expression in Prnp0/0 cells.(25) Remarkably, aldolase C, also implied in the glycolytic pathway, was recently identified as a novel interaction partner of PrPc.(26) The present results characterized a novel PrPc-dependent pathway, where silencing of PrPc inhibited glucose uptake by suppressing Glut1 expression. Also, our observation indicated the high correlation between PrPc, Glut1, and histological grades. Thus, PrPc would seem to be an ancillary protein required for the expression of glucose transporters.

An enhancement of glycolytic (anaerobic) metabolism and the resulting large increase in glucose requirement imply a need for a corresponding increase in glucose transport across the plasma membrane. The facilitated diffusion of glucose into cells is mediated by a family of homologous proteins, Glut1–5, that differ in their tissue distribution, affinities for glucose, and physiologic regulation.(27) Kinetic analysis of glucose transport revealed that inhibition of Glut1 by PrPc silencing abolished the Warburg effect in cancer cells, directly indicating that the greater degree of PrPc expression in the more malignant tumors most likely reflects a greater enhancement of glycolytic metabolism.

Glut1 has been shown to be regulated by HIF-1α, a master regulator of cellular and systemic responses to changes in available oxygen in the cellular environment.(28) In general, HIFs are overexpressed in multiple malignancies and crucial for the glycolytic phenotype of cancer cells.(29) We show here that HIF-2α, but not HIF-1α, is an important downstream target of PrPc explaining the mode of regulation of Glut1 expression at the molecular level in colorectal cancer cells. In immunohistological experiments we found the co-expression of Glut1 and HIF-2α in human colorectal carcinomas. Furthermore, a direct and specific binding of HIF-2α to the Glut1 promoter was detected, which was largely abolished by PrPc depletion. Similar to HIF-1α, continuous hypoxia leads to HIF-2α accumulation and the resulting transcriptional activation regulates several target genes that are also common to HIF-1.(30) As the inhibition of HIF-2α also reduced Glut1 expression and glucose transport capacity in colorectal cancer cells, the PrPc-HIF-2α-Glut1 axis is a relevant molecular mechanism that links PrPc expression to poor prognosis in malignancies depending on a high rate of glycolysis.

The attachment of PrPc to the plasma membrane through a glycosylphosphatidylinositol anchor may also be consistent with a role in cell-surface signaling or cell adhesion.(31) In the current context, the Fyn pathway lies downstream of PrPc, as PrPc silencing suppressed constitutive activation of Fyn, and vice versa. Moreover, inhibition of the Fyn pathway abolished the molecular events downstream of PrPc, suggesting that the Fyn pathway is required for PrPc-mediated HIF-2α transcriptional activity and expression of its target gene, Glut1.

Taken together, our results show for the first time that PrPc expression confers a direct significant growth advantage for colorectal carcinomas by a mechanism that involves the function of PrPc as an ancillary protein required for the function and expression of Glut1. Furthermore, our findings support the idea that the Warburg effect increases the malignant potential of colorectal cancers. Targeting pathways involved in glycolysis, such as the PrPc-Fyn-HIF2α-Glut1 axis, represents a promising avenue to inhibit the growth of colorectal cancer cells.

Disclosure Statement

The authors have no conflict of interest to declare.