The Anticancer Activity of Monosaccharides: Perspectives and Outlooks

Simple Summary

Despite recent advances in treatment options, such as chemotherapy, cancer continues to be the second-leading cause of death worldwide. Significant hurdles in the success of chemotherapy regimens include severe adverse effects, as well as drug resistance. As such, there is a strong requirement for novel pharmacological interventions in the treatment of cancer. Natural products, such as monosaccharides, are a promising potential treatment option for cancer due to their low toxicity, high solubility, and high specificity for tumour cells. Several naturally occurring and synthetically modified sugars have displayed toxicity in a variety of cancer and tumour cells.

Abstract

A major hallmark of cancer is the reprogramming of cellular metabolism from oxidative phosphorylation (OXPHOS) to glycolysis, a phenomenon known as the Warburg effect. To sustain high rates of glycolysis, cancer cells overexpress GLUT transporters and glycolytic enzymes, allowing for the enhanced uptake and consumption of glucose. The Warburg effect may be exploited in the treatment of cancer; certain epimers and derivatives of glucose can enter cancer cells and inhibit glycolytic enzymes, stunting metabolism and causing cell death. These include common dietary monosaccharides (ᴅ-mannose, ᴅ-galactose, ᴅ-glucosamine, ʟ-fucose), as well as some rare monosaccharides (xylitol, ᴅ-allose, ʟ-sorbose, ʟ-rhamnose). This article reviews the literature on these sugars in in vitro and in vivo models of cancer, discussing their mechanisms of cytotoxicity. In addition to this, the anticancer potential of some synthetically modified monosaccharides, such as 2-deoxy-ᴅ-glucose and its acetylated and halogenated derivatives, is reviewed. Further, this article reviews how certain monosaccharides can be used in combination with anticancer drugs to potentiate conventional chemotherapies and to help overcome chemoresistance. Finally, the limitations of administering two separate agents, a sugar and a chemotherapeutic drug, are discussed. The potential of the glycoconjugation of classical or repurposed chemotherapy drugs as a solution to these limitations is reviewed.

Keywords: monosaccharides, anticancer drugs, antitumour therapy, sugars, natural products, glycolysis

2.1. ᴅ-Mannose

ᴅ-mannose is an important dietary sugar that can be obtained from fruits such as oranges, peaches, and apples [15]. As the C-2 epimer of glucose, mannose plays important roles in the human body, for example, in the glycosylation of proteins [16]. Additionally, mannose is five times more active than glucose in non-enzymatic glycation [17]. Like glucose, mannose enters mammalian cells via the GLUTs, showing no preference or specificity for uptake by any of the 15 transporters [18]. Mannose is reported to possess immune regulatory functions and can suppress inflammatory disease [19].

Crucially, mannose has displayed notable anticancer properties. For example, in A549 cells, mannose has a strong inhibitory effect on proliferation, invasion, and metastasis [20]. Recently, Gonzalez et al. studied the effect of mannose on a variety of cancer cell lines, including KP-4 pancreatic cancer cells, and found that the monosaccharide significantly inhibited cell growth in vitro [21]. Mannose treatment at 25 mM more than halved the number of live KP-4 cells compared to the control. This cytostatic effect was confirmed in vivo, where mice bearing KP-4 xenografts were orally administered mannose, and this significantly impaired and retarded tumour growth. Together, these results suggest that mannose can inhibit cancer cell proliferation both in vitro and in vivo. It was hypothesised that mannose may interfere directly with glucose metabolism, but it was found that treatment of cancer cells with mannose had no effect on intracellular glucose levels. Instead, it was discovered that mannose accumulates in cancer cells as the glycolytic intermediate, mannose-6-phosphate (M6P), which suppresses phosphoglucose isomerase (PGI), hexokinase (HK), and other enzymes involved in the metabolism of glucose, hindering glycolysis. The inhibitory effect of mannose was found to be negatively correlated to the expression of phosphomannose isomerase (PMI), an enzyme that mobilises mannose towards glycolysis by converting M6P to fructose-6-phosphate (F6P). In colorectal tumour cells (RKO), which naturally express low levels of PMI, mannose significantly inhibited cell growth [21].

Saito et al. investigated the effect of mannose in leukaemia cells, which overexpress PMI [22]. Since leukaemia cells rapidly proliferate, an alternative nutrient source to glucose is often necessary for metabolism. Mannose was found to act as an energy source for metabolism in leukaemia cells, aiding glycolysis. However, when doses of mannose were used that exceeded the processing capacity of PMI, glycolysis and thus the growth of leukaemia cells were suppressed (Figure 3). These findings suggest that more evaluation is required for the use of mannose as a cancer therapy, and particular attention should be paid to the expression levels of PMI in the tumours of interest.

2.2. ᴅ-Glucosamine

ᴅ-glucosamine is an amino sugar that is made in the human body. Based on glucose, glucosamine is substituted at C-2 by an amine group. Glucosamine is involved in important biochemical processes, including in the synthesis of glycosylated proteins and lipids, as well as in the regulation of various signalling pathways [23,24]. An important medical use for glucosamine is in the treatment of osteoarthritis, owing to the powerful anti-inflammatory properties of this sugar [25]. Further, numerous cohort studies have found an association between habitual glucosamine supplementation and decreased cancer mortality [26,27].

The anticancer activity of glucosamine was first reported in 1953, when it was observed that in vivo administration of ᴅ-glucosamine hydrochloride to mice bearing Sarcoma 37 resulted in tumour growth retardation and the doubling of survival times of the animals [28]. In 1971, it was found that administration of glucosamine to rats bearing Walker 256 Carcinoma resulted in complete necrosis of the tumours within 40 h [29]. In 1980, Friedman and Skehan observed that treatment of C6 glioma cells with glucosamine results in cell lysis in vitro and this was potentiated by coculture with the membrane-active drug, lidocaine [30].

Despite the extensive study of glucosamine over the past 70 years, the exact mechanisms of the sugar’s anticancer activity are unclear. However, many studies have suggested that the cytotoxicity of glucosamine is related to its ability to inhibit protein synthesis and translation. For example, one study on the effect of glucosamine on DU145 and MDA-MB-231 cells found that the sugar dephosphorylated and thus inhibited the activity of the protein kinase, phosphorylated 70 ribosomal S6 protein-kinase (p70S6K), which is implicated in cell growth, proliferation, and cell cycle progression [31]. This inactivation resulted in growth inhibition in vitro, an observation that was selective to cancer cells. Similarly, Wang et al. observed an anti-proliferative effect of glucosamine (GlcN) on the human renal cancer cell lines, 786-O and Caki-1, in vitro [32]. This was attributed to glucosamine downregulating the cell cycle regulators, cyclin D1, cyclin-dependent kinase 4 (CDK4), and cyclin-dependent kinase 6 (CDK6). Meanwhile, glucosamine was found to upregulate the cell cycle inhibitors, p21 and p53. This resulted in cell cycle arrest in the phases G0/G1, as evidenced by the significantly higher volume of cells in the G1 phase compared to the S and G2 phases [32].

Glucosamine can also affect the activity of the forkhead box transcription factors (FOXOs), which modulate the expression of genes involved in apoptosis and cell cycle arrest [33,34]. Yu et al. demonstrated that glucosamine inhibited the phosphorylation of FOXO01 and FOXO03 in A549 cells, which interfered with their transcriptional activity and suppressed cellular proliferation [35]. The signal transducer and activator of transcription (STAT) transcription factors, which regulate oncogenic processes, such as proliferation and angiogenesis [36], are also affected by glucosamine. Chesnokov et al. demonstrated that glucosamine induced death in DU145 cells through the dephosphorylation of STAT3 and thus the suppression of this signalling pathway [37]. This resulted in cell cycle arrest at the G1 phase and apoptotic cell death.

A further way in which glucosamine causes an anticancer effect is through the induction of endoplasmic reticulum (ER) stress, which occurs when the ability of the ER to fold proteins becomes saturated [38]. Such stress promotes the unfolded protein response (UPR), a self-protective mechanism that shuts down protein translation and can also induce cell death via autophagy or apoptosis when stress is severe [39,40]. Hwang et al. observed reduced viability in glucosamine-treated U-87 MG cells, which was accompanied by an increase in the expression of markers of ER stress [41]. Treatment of U-87 MG cells with an autophagy inhibitor recovered cellular viability whilst treatment with an apoptosis inhibitor did not. Together, these data suggest that glucosamine induces autophagic death in U-87 MG cells via activating ER stress and triggering the UPR. Since many types of cancer are resistant to apoptosis due to mutations in the p53 tumour suppressor gene [42], targeting autophagy as an alternative cell death pathway is an important area in the development of chemotherapies [43].

Different mechanisms of action have been proposed to explain the anticancer effect of ᴅ-glucosamine; however, it remains unclear which mechanism is responsible. It is possible that a concatenation of several different effects may be attributed to the antineoplastic effects of glucosamine. Additionally, it is possible that the way in which glucosamine exerts its effects may be specific to the cancer cell line.

2.3. ᴅ-Galactose

ᴅ-galactose is the C-4 epimer of glucose and is one of the most abundant sugars in the human diet [44]. Galactose is widely involved in the biological function of human cells, particularly in the glycosylation of proteins and lipids, a process which plays a crucial role in cell–cell interaction, hormonal activity, immune modulation, and more [45,46,47]. In typical metabolism, galactose itself cannot undergo glycolysis so it enters the glycolysis pathway through its conversion into glucose-1-phosphate. This occurs in a series of steps known as the Leloir pathway [48,49]. However, when there is a large excess of galactose present in cells, alternative pathways can be employed in the metabolism of the sugar. One such pathway is the aldose reductase (AR) pathway, which metabolises galactose into the alcohol sugar, galactitol. Galactitol, poorly metabolised with poor liposolubility, cannot be transported across the cellular membrane. Instead, it accumulates within cells, increasing the osmotic pressure of the cells, which can result in oxidative stress and membrane rupture [50].

Exploiting the alternative metabolic pathways of galactose is one therapeutic target in anticancer drug development. Li et al. investigated the effects of galactose on the viability of some neuroblastoma cell lines (N2a, SH-SY5Y) [51]. At concentrations of 30 g/L or more, galactose inhibited the proliferation and suppressed the colony formation of neuroblastoma cells. Interestingly, galactose did not induce apoptotic cell death; instead, cell death was caused by necroptotic cell death, as evidenced by lactate dehydrogenase (LDH) leakage. Cotreatment of neuroblastoma cells with specific inhibitors of the AR pathway recovered the viability of the cancer cells, suggesting that this pathway is implicated in galactose-induced toxicity. It is possible that the leakage of LDH was a consequence of the accumulation of the AR metabolite, galactilol, leading to membrane rupture and cell death [51].

In another study, Shiratori et al. cultured A549 and HeLa cells in galactose and found that the glycolytic pathway was suppressed [52]. This metabolic suppression induced autophagy, but instead of promoting cell death, the autophagic pathway produced high levels of amino acids, shifting metabolism towards OXPHOS. This metabolic reprogramming helped in recovering the viability of the cells. It was suggested, therefore, that in some cancer cell lines, interfering with energy metabolism is not enough to cause cell death. Zheng et al. studied the effects of galactose on metabolism in breast cancer cell lines, whilst investigating whether particular oncogenes can affect the reprogramming of cellular metabolism [53]. Cells expressing AKT Serine/Threonine Kinase 1 (AKT1), an oncogene that may influence metabolism [54], were cultured in galactose-containing media, which resulted in rapid ROS-mediated cell death. This suggests that the AKT1-positive cells experienced oxidative stress and were unable to shift their metabolism from glycolysis to OXPHOS. However, after 15 days in galactose culture, the cells began to proliferate again, suggesting that the cells started to shift their metabolism and recover their viability. This study provides important evidence that certain oncogenes may expose metabolic vulnerabilities in particular cancer types. Additionally, it highlights that when developing therapies that target metabolism, the genetic profiles of tumours should be carefully considered.

2.4. ᴅ-Fructose

ᴅ-fructose is a simple sugar that is found in fruits and vegetables, as well as honey. Fructose is a structural isomer of glucose, and the two sugars can be converted into one another in the human body [55]. The metabolism of fructose is largely governed by fructokinase (FK), since this is the first enzyme that fructose encounters in its metabolic pathway. Fructose has a high affinity for GLUT5, and since many cancers overexpress this transporter, it is hypothesised that cancer cells may use fructose as an energy source [56,57,58].

Whilst the sugars discussed above could have potential in the fight against cancer, fructose is a far less attractive candidate. It has long been known that obesity and metabolic syndromes are associated with certain types of cancer [59] and various studies have suggested that consumption of high-fructose corn syrup (HFCS) may be responsible for the current global obesity and metabolic syndrome epidemics [60]. Furthermore, Goncalves et al. investigated the effect of high-fructose corn syrup on the growth of intestinal tumours in mice and found that consumption of just small amounts of fructose (around 3% of the total daily caloric intake of the animals) resulted in enhanced tumourigenesis [61].

In 2018, Weng et al. showed that incubation of non-small cell lung cancer, including A549 cells, with fructose resulted in enhanced proliferation of the cells [62]. The fructose-specific transporter, GLUT5, is encoded for by solute carrier family 2 member 5 (SLC2A5) [63]. Overexpression of SLC2A5, and hence GLUT5, is associated with enhanced tumourigenesis, migration, and invasion, all of which contribute to cancer metastasis [62]. In a study with MDA-MB-468 breast cancer cells, which also overexpress GLUT5 [64], fructose induced an aggressive phenotype in the cells that was associated with enhanced migratory and invasive characteristics [65]. Together, these results show that fructose is not an ideal candidate in the development of anticancer agents since it can contribute to tumour progression and metastasis.

2.5. ʟ-Fucose

ʟ-fucose is a dietary 6-deoxy sugar that plays a major role in mammalian biology. Unlike many of the standard hexose sugars found in the human body, fucose is mainly present in the ʟ configuration. The process of attaching a fucose unit to a molecule is known as fucosylation. Fucosylated glycans, synthesised by fucosyltransferases (FUTs), are vastly important in regulating inflammatory responses, cell signalling, transcription, protein folding, and cell–cell adhesion [66]. In cancer, fucose metabolism is altered, which can result in the aberrant expression of proteins. The potential consequences of this are evasion from immune control, invasion, metastasis, as well as reduced adhesion, and more [67,68,69]. In fact, increased fucosylation has been associated with the acquisition of metastatic properties [70,71,72]. At the same time, fucosylation is reported to reduce metastatic properties in melanoma cells [72], head and neck cancer cells [73], and hepatocellular carcinoma cells [74]. Additionally, exogenous fucose was found to rescue fucosylation in Caco-2 cells and to increase the efficiency of immunotherapy via upregulating fucosylation [75,76]. The differences in these results may be explained by the different types of tumour cells investigated in each study. Clearly, the exact extent of the protumour or antitumour effects of fucosylation warrants further exploration. As such, some researchers have investigated the effects of exogenous fucose in different models of cancer.

The effect of ʟ-fucose on solid rat mammary adenocarcinomas was first explored in a series of studies by Rosato et al. In vitro administration of fucose on mammary tumour cells resulted in significant growth inhibition in a dose-dependent manner, accompanied by morphological changes to the cells. In vivo studies showed significant reductions in the diameters of rat mammary tumours following daily IP infusion of fucose for 10 days. These early studies indicated the potential of fucose as a powerful anticancer and antitumour agent [77,78,79].

Tomsik et al. investigated the effect of in vivo-administered fucose on the growth, mitotic activity, and metastatic activity of solid Ehrlich carcinomas in immunocompetent mice [80]. IP administration of fucose significantly inhibited the growth of tumours, with a dose of 5 g/kg/day resulting in a 47% growth inhibition after 11 days. Despite this, fucose treatment displayed only a slight improvement on the survival of the tumour-bearing mice. Histological analysis of the treated tumours revealed significant decreases in mitotic indices, suggesting that fucose caused a reduction in the number of dividing tumour cells. These data suggest that fucose could be a safe and effective anticancer agent.

Recently, Yao et al. investigated whether the anticancer effect of fucose is a result of the sugar upregulating the fucosylation of proteins in cancer cells [81]. As such, the effect of ʟ-fucose on the colorectal cancer (CRC) cell line, HCT-116, a cancer accompanied with aberrant glycosylation, was studied [82]. Fucose supplementation significantly decreased the proliferation rate and the migration of HCT-116 cells, but not HCoEpic cells (healthy colonic) or HT-29 cells (CRC cells), suggesting cell-type specificity. Furthermore, fucose treatment increased the levels of fucosylation in HCT-116 cells only. Mechanistic studies revealed that fucose supplementation in HCT-116 cells upregulated the expression of genes related to serine biosynthesis, such as phosphoserine aminotransferase (PSAT1). Knocking down of this gene resulted in decreased intracellular serine concentrations, diminished fucosylation, and an impaired ability of fucose to inhibit cell migration and proliferation. Concurrently, ʟ-fucose treatment increased the intracellular concentration of serine in HCT-116 cells. This suggests that exogenous ʟ-fucose may promote fucosylation in colorectal cancer by enhancing the intracellular accumulation of serine, an important substrate in O-glycosylation [83], via the upregulation of PSAT1.

3.1. Xylitol

Xylitol is a sugar alcohol that occurs naturally in numerous plants, including fruits and berries, mushrooms, corn husks, and the birch tree. Xylitol can also be obtained from the enzymatic or catalytic hydrogenation of ᴅ-xylose, an aldopentose [86]. The structures of these two related compounds are displayed in Figure 4. Commonly used as an artificial sweetener, xylitol is popular with diabetics and those on a low-carbohydrate diet since the sugar has a low glycaemic index and induces a low insulinaemic response [87]. In addition to being a sugar replacer, xylitol is commonly used in oral health care due to its potent antibacterial and anticariogenic properties [88]. Interestingly, xylitol is reported to suppress anaerobic glycolysis in bacteria [89]. Despite being of plant origin, xylitol can be used in mammalian glycolysis [90]. However, unlike glucose, the plant sugar generates ATP via the pentose phosphate pathway (PPP), and this is achieved through the conversion of xylitol to xylulose via xylitol dehydrogenase [91]. Due to being highly water soluble and displaying no toxicity or carcinogenicity in humans, xylitol is an attractive potential anticancer drug.

In 2014, Park et al. reported that xylitol inhibited the in vitro proliferation of a variety of human cancer cell lines in a dose-dependent manner, with the most pronounced toxic effect observed in A549 lung cancer cells [92]. The IC₅₀ of xylitol in cancer cells was significantly lower than that of xylitol in healthy fibroblastic cells, suggesting some selectivity of xylitol towards cancer cells. This could be due to an overexpression of xylitol transporters in cancer cells compared to healthy cells. Mechanistic studies revealed that xylitol treatment did not induce apoptosis in A549 cells. However, when A549 cells were cotreated with the autophagy inhibitor, 3-methyladenine, the viability of the xylitol-treated cells was recovered. This suggests that the toxicity of xylitol in A549 cells is caused by the induction of autophagy [92].

More recent studies with xylitol have shown its ability to suppress glycolysis. Trachootham et al. partially substituted glucose for xylitol in the culture media of CAL-27 oral cancer cells and found that this significantly suppressed their proliferation and ATP levels [93]. Further, xylitol treatment significantly suppressed the activity of the enzyme, phosphofructokinase (PFK), a key regulator of glucose metabolism. This suggests that the sugar interfered with the glycolytic activity of the cells. This effect was not observed in healthy oral keratinocytes, suggesting a selectivity of xylitol for cancer cells. A follow-up study investigated whether the antiglycolytic effect of xylitol could be observed in vivo [94]. To achieve this, mice bearing CAL-27 xenografts were administered a glucose and xylitol-containing feed. This treatment resulted in significant retardation of tumour proliferation; however, there were no significant reductions in the volumes of the tumours. Mechanistic studies revealed that the xylitol-containing feed significantly reduced the expression of PFK, and this was accompanied by a non-significant reduction in the activity of the enzyme. This suggests that xylitol may have some effect on glycolysis in vivo.

Tomonobu et al. investigated the effects of xylitol treatment on MeWo melanoma cells in vitro and found that the sugar induced apoptosis in a dose-dependent manner [95]. Apoptosis was followed by a marked elevation in ROS and a significant decrease in the levels of reduced glutathione, an antioxidant that is involved in DNA repair. ROS were induced in cancer cell lines but not in healthy cells, suggesting a specificity of xylitol for cancer. Mechanistically, xylitol-mediated apoptosis was a result of the upregulation of CHAC1, an enzyme associated with ER stress and apoptosis, which is responsible for the degradation of glutathione [96]. In vivo, xylitol was administered to mice bearing MeWo xenografts [95]. Oral administration of xylitol had no effect on tumour volume, which was likely caused by acid-mediated inactivation of the sugar in the gut. This may also explain why in the study discussed above [94], oral administration of xylitol caused no significant reductions in tumour volumes. However, IV administration of xylitol resulted in attenuation of the tumour growth rate and significant reductions in tumour masses and volumes [95]. Furthermore, the tumour fronts of xylitol-treated xenografts were smooth, suggesting that the sugar can reduce the invasiveness of cancer cells. Mass spectroscopic analysis of tumours indicated no apparent changes in the major glycolytic metabolites, suggesting that xylitol did not affect glycolysis in vivo. However, when xylitol was co-administered with tauroursodeoxycholic acid (TUDCA), an inhibitor of ER stress, tumour suppression was inhibited, demonstrating that ER stress may be an anticancer mechanism of the sugar.

3.2. ᴅ-Allose

ᴅ-allose is the C-3 epimer of glucose and is rarely encountered in nature. This rare sugar is non-toxic, odourless, and highly water soluble [97]. Allose has been isolated from numerous natural resources, including potato leaves [98] and Indian seagrasses [99]. Chemical syntheses have also been employed in the production of allose; however, these are often complex reactions, with poor selectivity and unwanted side-products. Thus, enzymatic syntheses are more widely used to produce this rare sugar [100].

Despite the rarity of allose, a few anticancer studies have been conducted with the sugar. For example, allose has suppressed the in vitro proliferation of cervical and skin cancer cells [101], ovarian cancer cells [102], and prostate cancer cells [103]. Hirata et al. reported a dose-dependent inhibitory effect of allose on leukaemia cell proliferation [104]. This effect was most pronounced in the cell line, MOLT-4F. Treatment of leukaemia cells with allose resulted in cell cycle arrest, specifically in the G1 phase. This was accompanied by significantly upregulated expression levels of thioredoxin-interacting protein (TXNIP), a gene that has been reported to be a tumour [105] and metastasis suppressor [106]. Further, allose treatment significantly upregulated the levels of p27^kip1, a cell cycle inhibitor [107]. At the same time, allose treatment did not cause apoptotic cell death, suggesting that allose does not affect the expression of proteins and genes associated with apoptosis. TXNIP is also known to regulate glucose metabolism in skeletal muscle and adipocytes [108]. Thus, allose treatment resulted in a significant decrease in the uptake of glucose into MOLT-4F cells, suggesting that the rare sugar can inhibit the metabolic activity of the cells.

Hoshikawa et al. also examined the effects of allose on TXNIP expression in cancer [109]. Allose treatment significantly inhibited the proliferation of two human oral squamous cancer cell lines in vitro (HSC-3 and Ca9-22). TXNIP expression levels were markedly increased in both cell lines, suggesting that TXNIP gene expression plays a role in suppressing cancer cell growth. This overexpression was also measured in tumour tissue following in vivo studies, in which mice bearing HSC-3 xenografts were intravenously administered a solution of allose. This resulted in a reduction in tumour volume of 39% compared to the control. These data suggest that allose, as well as other agents that can enhance the activity of TXNIP, are attractive candidates in the treatment of cancer. In addition, it was found that allose treatment significantly downregulated the expression of GLUT1 in some human cancer cell lines (HuH-7, MDA-MB-231, and SH-SY5Y) [110]. As evidenced by 2-DG uptake assays, a reduction in the expression of GLUT1 resulted in significant decreases in glucose uptake into cancer cells [110]. Together, these data suggest that allose can impart significant effects on protein expression in cancer cells, and that its consequential deleterious effect on glucose uptake is a mechanism by which the rare sugar exerts its anticancer effects.

Allose is also known to affect the expression of apoptotic proteins. Naha et al. reported an inhibitory effect of allose on the proliferation of human prostate cancer cells (DU145) [111]. Meanwhile, the rare sugar had no remarkable effect on the viability of healthy human prostate epithelial cells, suggesting a specificity of allose for cancer. Prostate homeostasis in man exists as an equilibrium between cell proliferation by mitosis and cell death through apoptosis. These activities are regulated by the antiapoptotic protein, B-cell leukaemia/lymphoma 2 protein (Bcl-2), and the pro-apoptotic protein, Bcl-2-associated X protein (Bax), respectively [112]. Allose treatment significantly decreased the expression of Bcl-2 in prostate cancer cells whilst concomitantly increasing the expression of Bax, in a dose-dependent manner. In line with this, allose caused significant apoptotic cell death in prostate cancer cells. Cell cycle analysis revealed that allose triggered the significant arrest of cells in the G0/G1 phase of the cell cycle. This is expected since cell cycle regulation and apoptosis are closely related. However, this result agrees only partly with the work discussed above by Hirata et al. [104], who showed that allose caused cell cycle arrest in the G1 phase; however, allose did not cause apoptotic cell death. These differences may be explained by the different types of cancer cell lines used.

3.3. ʟ-Sorbose

ʟ-sorbose is a rare natural sugar that is the C-3 epimer of ᴅ-fructose. Due to its structural similarity to fructose, sorbose is internalised into cells mainly by the fructose-specific GLUT5 transporters [113]. The presence of sorbose in foods and other natural sources is not well known; however, the sugar can be produced from the bacterial-mediated dehydrogenation of ᴅ-sorbitol, which is found in numerous fruits [114]. Sorbose is an important starting material in the manufacture of vitamin C, as well as in the synthesis of other rare sugars, such as ʟ-tagatose [114]. Research into the use of sorbose as a therapeutic agent has been largely limited to antimicrobial [115] and antifungal studies [116], as well as studies concerning glycaemic control [117].

However, a recent landmark paper unveiled the powerful anticancer properties of the sugar [118]. Sorbose was reported to impart a significant inhibitory effect on the viability of various human cancer cell lines (Huh-7, HepG2, A549, HeLa, MCF-7, and K562). Mechanistically, sorbose increased the levels of cleaved caspase 3 and the ratio of BAX/Bcl2, suggesting that the sugar can induce apoptosis in cancer cells. It was found that once inside cells, sorbose is converted into sorbose-1-phosphate (S1P) by ketohexokinase (KHK). S1P subsequently inactivates the glycolytic enzyme, HK, hindering glycolysis, inducing ROS production, and, consequently, triggering apoptosis. Additionally, S1P was found to downregulate the expression of KHK-A, an isoform of KHK that mediates the phosphorylation of proteins including p62, a selective autophagy receptor. Phosphorylation of p62 by KHK-A in regular cell metabolism results in the activation of the transcription factor, erythroid 2-related factor 2 (Nrf2), which induces ROS-fighting antioxidant genes [119]. Inactivation of KHK-A by sorbose treatment caused the attenuation of the Nrf2 pathway, thus reducing the antioxidant defence system in the cancer cells. These two inhibitory effects of S1P were found to work in synergy to induce apoptosis in cancer cells. Mice bearing Huh-7 xenografts were orally administered sorbose, which resulted in significant tumour growth inhibition and reductions in tumour volumes. Following sorbose treatment, no notable weight losses were observed in the animals and the plasma glucose and insulin levels were unchanged, showing that the sugar does not affect glycaemic indices in mice. These observations suggest that sorbose could be used safely as an anticancer drug.

3.4. ʟ-Rhamnose

ʟ-rhamnose is a natural 6-deoxy methyl pentose sugar that is found in plants and bacteria, but is not made in mammals [120]. Hence, this sugar is recognised in the human body as a xenoantigen. Interestingly, human serum contains a high concentration of anti-ʟ-rhamnose antibodies; thus, exposing malignant cells to rhamnose can induce an antitumour immune response in vivo [121]. Despite its non-mammalian origin, rhamnose can serve as a substrate in fucose metabolism and can be incorporated into glycosides in the human body [122]. As such, it can be hypothesised that rhamnose could negatively influence fucosylation and thus the behaviour of malignant cells. These two potential mechanisms of toxicity make rhamnose an attractive agent in anticancer drug development.

In 2014, Tomsik et al. evaluated the in vitro antitumour effect of rhamnose in Ehrlich ascites tumour cells and HL-60 leukaemia cells [122]. Surprisingly, rhamnose treatment at 20 mM caused no significant reductions in the proliferation of either cell type. Rhamnose-functionalised conjugates have also been explored for the targeted destruction of tumour cells. For example, Li et al. developed a targeted rhamnose-functionalised liposome (Rha-lip) and found it to be active against various types of cancer cells [123]. Incubation of HeLa and A549 cells with Rha-lip and human serum, a source of anti-rhamnose antibodies, resulted in significant in vitro toxicity. This suggests that for rhamnose-induced cytotoxicity to be achieved, malignant cells must be cultured in the presence of anti-rhamnose antibodies. Additionally, mice bearing 4T1 breast cancer tumours were treated with Rha-lip via IV injection, which significantly retarded tumour growth. This was accompanied by no significant weight losses in the mice treated with Rha-lip, suggesting that rhamnose-functionalised liposomes could be safe and effective antitumour agents [124].

In a different study, Tomsik et al. administered rhamnose to solid Ehrlich tumour-bearing mice via IP injection [80]. Rhamnose treatment at a dose of 3 g/kg/day resulted in significant tumour growth inhibition. Additionally, this dose of the sugar significantly prolongated the lives of the animals. Further, there was a significant decrease in the mitotic indices of the tumours following rhamnose treatment, as well as evidence of necrotic processes. These data suggest that rhamnose possesses antitumour properties. Following ex vivo analysis of the organs, rhamnose revealed no metastases, suggesting that the sugar might help to prevent tumour invasion. Interestingly, the mice used in this study were not immunised with anti-rhamnose antibodies. This suggests (1) the strain of mice used in this study (NMRI) may have naturally higher basal levels of anti-rhamnose antibodies than BALB/c mice and/or (2) the mechanism of toxicity exerted by rhamnose in this study was not related to immunomodulation. It was also concluded that rhamnose does not interfere with energy metabolism like other monosaccharides can [122]. Since rhamnose is recognised by enzymes involved in fucosylation, it is possible that rhamnose exerts its anticancer effect via interfering with fucose metabolism. However, much more research is required to elucidate the mechanism of action of ʟ-rhamnose.

The effects of common and rare monosaccharides in in vitro and in vivo models of cancer, along with their proposed mechanisms of action, are summarized in Table 1.

4.1. 2-Deoxy-ᴅ-glucose

2-Deoxy-ᴅ-glucose (2-DG) (Figure 5) is a synthetic, non-metabolisable glucose analogue; the hydroxyl group on carbon-2 is replaced with a hydrogen. Due to its structural similarity to glucose, 2-DG competes with glucose in the first stage of metabolism, thus inhibiting cellular ATP production [125]. 2-DG shows activity against viruses, including COVID-19, and efficacy in models of polycystic kidney disease, epilepsy, and importantly, in cancer [126]. Despite its synthetic origin, the 2-DG moiety exists in a number of natural products, including in anthracyclines and erythromycins, which may explain its vast biological applicability. Like glucose, 2-DG enters cells via the GLUT transporters and is phosphorylated to 2-deoxy-ᴅ-glucose-6-phosphate (2DG-6P) by HK [127]. However, unlike glucose, 2DG-6P cannot continue down the glycolytic pathway because it is not a substrate for PGI or glucose-6-phosphate dehydrogenase, and so accumulates in cells [127]. As such, the partial metabolism of 2-DG consumes ATP but does not contribute to the energy payoff since glycolysis cannot be completed. Studies have shown that 2-DG also affects the metabolism of glucose through competitively inhibiting PGI and so preventing the conversion of glucose-6-phosphate to fructose-6-phosphate [128], as well as non-competitively inhibiting HK [129], thus preventing the phosphorylation of glucose. This catabolic block contributes to the depletion of cellular energy.

In cancer, the inhibition of glycolysis by 2-DG is more pronounced under hypoxia. This is because, under normoxic conditions, cancer cells can flexibly shift their metabolism. However, under hypoxic conditions, cancer cells rely solely on glycolysis. In 2002, Liu et al. showed how the ATP-reducing effects of 2-DG are significantly potentiated under hypoxic conditions [130]. MIA-PaCa-2 cells were treated with 2-DG under hypoxic (<0.1% O₂) or normoxic (21% O₂) conditions. At a dose of 24 mM, 2-DG treatment caused ATP levels in the normoxic cells to reduce to around 60% compared to the control, whilst treatment with the same concentration of 2-DG under hypoxic conditions resulted in a reduction in ATP levels to less than 10%. These findings suggest that 2-DG is a more potent glycolytic inhibitor under hypoxic conditions. Interestingly, under hypoxia, 2-DG inhibits autophagy, which may be due to the severe depletion in ATP levels under these conditions. This may be an advantageous result since autophagy can be a pro-survival mechanism in cancer [131].

Interestingly, 2-DG can exert significant toxicity to tumour cells under normoxic conditions, suggesting that a mechanism other than glycolytic inhibition is also at play [132]. A clue to this mechanism comes from the fact that 2-DG is not only structurally similar to glucose, but it is also similar to mannose, since glucose and mannose are C-2 epimers of one another. In the 1970s, Shwartz et al. demonstrated that in certain eukaryotic cells, 2-DG inhibits N-linked glycosylation [133,134,135,136]. The authors showed how the metabolite 2-DG-GDP competes with mannose-GDP for incorporation onto N-acetyl glucosamine residues, preventing the assembly of lipid-linked oligosaccharides (LLOs). Intriguingly, the addition of exogenous mannose to 2-DG-treated tumour cells recovers their viability, which suggests that the inhibition of protein glycosylation is a major mechanism of toxicity of 2-DG [137]. Kurtoglu et al. found that inhibiting N-linked glycosylation with 2-DG results in disruption in the folding of important glycoproteins, leading to a build-up of unfolded proteins in the ER [132]. This results in the UPR, which can lead to the activation of ER stress-specific apoptotic pathways.

The UPR and autophagy are also linked, which may explain why 2-DG has been reported to induce autophagy in vitro [138,139] and in patients [138]. Xi and colleagues found that ER stress induced by 2-DG triggers calcium-mediated activation of AMP-activated protein kinase (AMPK), which leads to the stimulation of autophagy [140]. This suggests a role of AMPK as a sensor of ER stress. In addition to this, 2-DG is known to activate AMPK and induce death receptor-mediated apoptosis. Pradelli et al. observed that AMPK activation in human cancer cell lines (HeLa and U-937) is caused by glycolytic inhibition and the subsequent depletion in ATP levels [141]. AMPK activation was found to lead to the inhibition of the mammalian target of rapamycin (mTOR), a downstream target of AMPK, which can promote tumour growth [142]. Blocking this pathway resulted in the downregulation of Mcl-1, a protein of the antiapoptotic Bcl-2 family, which caused an enhancement in apoptosis. Clearly, AMPK is an important mediator of cell death and the pathway by which cancer cells take to induce death (via autophagy or apoptosis) may be cell line-dependent.

The PPP is a metabolic pathway parallel to glycolysis that regenerates nicotinamide adenine dinucleotide phosphate hydrogen (NADPH), a H₂O₂ scavenger, from nicotinamide adenine dinucleotide phosphate (NADP⁺) [143]. Coleman et al. treated PaCa-2 cells with 2-DG and observed depleted NADPH levels, which led to the decreased production of reduced glutathione, a major antioxidant [144]. These data suggest that 2-DG can inhibit the PPP, in addition to blocking glycolysis. Thus, 2-DG can be described as enhancing ROS levels in cancer cells in a twofold fashion: (1) by inhibiting glycolysis and reducing the levels of pyruvate, also an H₂O₂ scavenger, and (2) by blocking the PPP and depleting NADPH levels.

In summary, the cell-killing effect of 2-DG appears to follow a hierarchy; at low doses and under normoxia, N-linked glycosylation is inhibited, which results in ER stress and the UPR. At moderate doses of 2-DG, glycolysis is inhibited and finally, at high doses, 2-DG treatment results in the blockage of the PPP [145]. A schematic summarising the effects of 2-DG in cancer cells is presented in Figure 6. The use of 2-DG as a monotherapy in humans may be limited since higher doses of 2-DG have been linked to insulin-induced hypoglycaemic symptoms, such as flushing and sweating, during clinical trials [146]. 2-DG may be most effective when structurally modified or used in conjunction with a traditional chemotherapy, as we shall discuss in the next sections.

4.2. Halogenated 2-DG

The glucose analogue, ¹⁸FDG, was first developed in 1976 as a radiotracer to study and map glucose metabolism in the brain via positron emission tomography (PET) imaging [147]. Taking advantage of the Warburg effect and the increased demand of cancer cells for glucose, ¹⁸FDG and PET are now used widely as tools for localising and diagnosing metabolically active tumours. ¹⁸FDG differs from 2-DG in that it has an ¹⁸fluorine substituent on C-2, in place of the H atom. The choice to position the radiolabelled fluorine atom on C-2 is significant since substitution at this carbon does not affect the ability of the sugar to traverse the blood–brain barrier (BBB) [147].

Structurally, ¹⁸FDG is more similar to glucose than 2-DG. This is because the fluorine atom at C-2 is energetically and conformationally more similar to the hydroxyl group of the glucose molecule [9]. As such, ¹⁸FDG and its non-radiolabelled counterpart, 2-FDG, are more potent glycolytic inhibitors than 2-DG, suggesting that 2-FDG may be a better substrate for HK. Molecular modelling studies by Lampidis et al. revealed that the binding energy of the metabolite, 2-FDG-6P, for the allosteric site of HK was lower than that of 2-DG-6-P, which suggests that 2-FDG is the better non-competitive inhibitor of HK and thus glycolysis [148]. This eventuality is compounded by the fact that in experimental studies, 2-FDG more potently killed hypoxic osteosarcoma cells than 2-DG did, at the same dose. Furthermore, cells incubated with 2-FDG more potently reduced lactate levels than those incubated with 2-DG, suggesting that 2-FDG is a more powerful inhibitor of glycolysis [148]. As well as investigating the anticancer potential of the fluorinated deoxyglucose analogue, Lampidis and colleagues developed 2-chloro-2-deoxy-ᴅ-glucose (2-CDG) and 2-bromo-2-deoxy-ᴅ-glucose (2-BDG) [148]. Studies performed on the series of halogenated deoxyglucose analogues (2-XDGs) revealed that 2-FDG was the most effective at inducing cytotoxicity, followed by 2-CDG then 2-BDG. Accompanying this, 2-FDG was the best at reducing intracellular lactate levels, suggesting that 2-FDG is the most potent inhibitor of glycolysis. It appears that the cytotoxic potential of the halogenated 2-DG analogues decreases with the increasing size of the halogen substituent (Br > Cl > F). This is likely due to the decreasing ability of the 2-XDGs to effectively bind to HK as the size of the C-2 substituent increases, leading to decreased glycolytic inhibition and decreased cell killing [148]. The structures of some halogenated deoxyglucose compounds are displayed in Figure 5.

Despite the ability of 2-FDG to powerfully inhibit glycolysis under hypoxic conditions, there is little to no antiglycolytic effect and thus cell killing of the sugar when exposed to cells growing under normoxia [148]. This is because, as previously discussed, when cancer cells are grown under normoxia, they can rely on forms of metabolism other than glycolysis. One of the main mechanisms of toxicity of 2-DG is its ability to compete with mannose due to their structural similarities and inhibit protein glycosylation [132], leading to ER stress and the UPR. However, since 2-FDG has a fluorine substituent on its carbon 2 in place of a H atom, it is no longer structurally analogous to 2-deoxy-ᴅ-mannose, so has no deleterious effect on protein glycosylation. This means that it cannot exert toxicity to cells via interfering with protein glycosylation. As such, 2-FDG is a less effective anticancer agent than 2-DG.

4.3. WP1122—O-Acetylated 2-DG

An O-acetylated version of 2-DG, WP1122, was developed by Dr. Priebe and colleagues to enhance the efficacy of the deoxy sugar in models of glioblastoma (GBM) [149]. WP1122, also known as 3,6-di-O-acetyl-2-deoxy-ᴅ-glucose, is substituted at carbons 3 and 6 with acetoxy groups in place of hydroxyl groups (Figure 5). This chemical modification renders the sugar more non-polar, allowing it to cross the BBB via passive diffusion, as opposed to relying on uptake by the GLUT transporters [9]. Once inside cells, WP1122 undergoes deacetylation by intracellular esterases, reforming 2-DG and commencing inhibition of the glycolytic pathway. Thus, WP1122 essentially acts as a prodrug of 2-DG. Preclinical in vivo studies performed by Priebe et al. revealed that when WP1122 is delivered orally, plasma concentrations of 2-DG are twofold higher than when pure 2-DG alone is administered. Furthermore, WP1122 is well tolerated by animals and was able to significantly prolong the life of GBM-bearing mice [149]. Taken together with in vitro data that demonstrate the ability of WP1122 to effectively inhibit glycolysis under hypoxic and normoxic conditions in U-87 MG cells [149], the prodrug has excellent potential as an anticancer and antitumour agent. WP1122 is currently undergoing phase I clinical trials.

Table 2.

Some examples of efficacious preclinical studies combining anticancer therapies with different monosaccharides.

• 1.Sung H., Ferlay J., Siegel R.L., Laversanne M., Soerjomataram I., Jemal A., Bray F., Bsc M.F.B., Me J.F., Soerjomataram M.I., et al. Global Cancer Statistics 2020: GLOBOCAN Estimates of Incidence and Mortality Worldwide for 36 Cancers in 185 Countries. CA Cancer J. Clin. 2021;71:209–249. doi: 10.3322/caac.21660. [DOI] [PubMed] [Google Scholar]
• 2.Hanahan D., Robert A. Weinberg, Hallmarks of Cancer: The Next Generation. Cell. 2011;144:646–674. doi: 10.1016/j.cell.2011.02.013. [DOI] [PubMed] [Google Scholar]
• 3.Zheng H.C. The molecular mechanisms of chemoresistance in cancers. Oncotarget. 2017;8:59950–59964. doi: 10.18632/oncotarget.19048. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 4.Nurgali K., Jagoe R.T., Abalo R. Editorial: Adverse Effects of Cancer Chemotherapy: Anything New to Improve Tolerance and Reduce Sequelae? Front. Pharmacol. 2018;9:245. doi: 10.3389/fphar.2018.00245. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 5.Ortega A.D., Sánchez-Aragó M., Giner-Sánchez D., Sánchez-Cenizo L., Willers I., Cuezva J.M. Glucose avidity of carcinomas. Cancer Lett. 2009;276:125–135. doi: 10.1016/j.canlet.2008.08.007. [DOI] [PubMed] [Google Scholar]
• 6.Cunha A., Rocha A.C., Barbosa F., Baião A., Silva P., Sarmento B., Queirós O. Glycolytic Inhibitors Potentiated the Activity of Paclitaxel and Their Nanoencapsulation Increased Their Delivery in a Lung Cancer Model. Pharmaceutics. 2022;14:2021. doi: 10.3390/pharmaceutics14102021. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 7.Nowak N., Kulma A., Gutowicz J. Up-regulation of Key Glycolysis Proteins in Cancer Development. Open Life Sci. 2018;13:569–581. doi: 10.1515/biol-2018-0068. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 8.Warburg O., Wind F., Negelein E. The Metabolism of Tumors in the Body. J. Gen. Physiol. 1927;8:519–530. doi: 10.1085/jgp.8.6.519. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 9.Pajak B., Siwiak E., Sołtyka M., Priebe A., Zieliński R., Fokt I., Ziemniak M., Jaśkiewicz A., Borowski R., Domoradzki T., et al. 2-Deoxy-d-Glucose and Its Analogs: From Diagnostic to Therapeutic Agents. Int. J. Mol. Sci. 2019;21:234. doi: 10.3390/ijms21010234. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 10.Hossain F., Andreana P.R. Developments in Carbohydrate-Based Cancer Therapeutics. Pharmaceuticals. 2019;12:84. doi: 10.3390/ph12020084. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 11.Zhao Y., Dunmall L.S.C., Cheng Z., Wang Y., Si L. Natural products targeting glycolysis in cancer. Front. Pharmacol. 2022;13:1036502. doi: 10.3389/fphar.2022.1036502. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 12.Barchi J.J., Jr. Emerging roles of carbohydrates and glycomimetics in anticancer drug design. Curr. Pharm. Des. 2000;6:485–501. doi: 10.2174/1381612003400876. [DOI] [PubMed] [Google Scholar]
• 13.Zhao M., Wei F., Sun G., Wen Y., Xiang J., Su F., Zhan L., Nian Q., Chen Y., Zeng J. Natural compounds targeting glycolysis as promising therapeutics for gastric cancer: A review. Front. Pharmacol. 2022;13:1004383. doi: 10.3389/fphar.2022.1004383. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 14.Stylianopoulos C.L. Encyclopedia of Human Nutrition. 2nd ed. Elsevier; San Diego, CA, USA: 2005. Carbohydrates: Chemistry and Classification; pp. 303–309. [Google Scholar]
• 15.Herman R.H. Mannose metabolism. I. Am. J. Clin. Nutr. 1971;24:488–498. doi: 10.1093/ajcn/24.4.488. [DOI] [PubMed] [Google Scholar]
• 16.Alton G., Hasilik M., Niehues R., Panneerselvam K., Etchison J.R., Fana F., Freeze H.H. Direct utilization of mannose for mammalian glycoprotein biosynthesis. Glycobiology. 1998;8:285–295. doi: 10.1093/glycob/8.3.285. [DOI] [PubMed] [Google Scholar]
• 17.Bunn H.F., Higgins P.J. Reaction of Monosaccharides with Proteins: Possible Evolutionary Significance. Science. 1981;213:222–224. doi: 10.1126/science.12192669. [DOI] [PubMed] [Google Scholar]
• 18.Mueckler M., Thorens B. The SLC2 (GLUT) family of membrane transporters. Mol. Aspects Med. 2013;34:121–138. doi: 10.1016/j.mam.2012.07.001. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 19.Zhang W., Cheng H., Gui Y., Zhan Q., Li S., Qiao W., Tong A. Mannose Treatment: A Promising Novel Strategy to Suppress Inflammation. Front. Immunol. 2021;12:756920. doi: 10.3389/fimmu.2021.756920. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 20.Wang Y., Xie S., He B. Mannose shows antitumour properties against lung cancer via inhibiting proliferation, promoting cisplatin-mediated apoptosis and reducing metastasis. Mol. Med. Rep. 2020;22:2957–2965. doi: 10.3892/mmr.2020.11354. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 21.Gonzalez P.S., O’prey J., Cardaci S., Barthet V.J.A., Sakamaki J.-I., Beaumatin F., Roseweir A., Gay D.M., Mackay G., Malviya G., et al. Mannose impairs tumour growth and enhances chemotherapy. Nature. 2018;563:719–723. doi: 10.1038/s41586-018-0729-3. [DOI] [PubMed] [Google Scholar]
• 22.Saito Y., Kinoshita M., Yamada A., Kawano S., Liu H., Kamimura S., Nakagawa M., Nagasawa S., Taguchi T., Yamada S., et al. Mannose and phosphomannose isomerase regulate energy metabolism under glucose starvation in leukemia. Cancer Sci. 2021;112:4944–4956. doi: 10.1111/cas.15138. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 23.Dalirfardouei R., Karimi G., Jamialahmadi K. Molecular mechanisms and biomedical applications of glucosamine as a potential multifunctional therapeutic agent. Life Sci. 2016;152:21–29. doi: 10.1016/j.lfs.2016.03.028. [DOI] [PubMed] [Google Scholar]
• 24.Konopka J.B. N-acetylglucosamine (GlcNAc) functions in cell signaling. Scientifica. 2012;2012:489208. doi: 10.6064/2012/489208. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 25.Reginster J.-Y., Neuprez A., Lecart M.-P., Sarlet N., Bruyere O. Role of glucosamine in the treatment for osteoarthritis. Rheumatol. Int. 2012;32:2959–2967. doi: 10.1007/s00296-012-2416-2. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 26.Zhou J., Wu Z., Lin Z., Wang W., Wan R., Liu T. Association between glucosamine use and cancer mortality: A large prospective cohort study. Front. Nutr. 2022;9:947818. doi: 10.3389/fnut.2022.947818. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 27.Li G., Zhang X., Liu Y., Zhang J., Li L., Huang X., Thabane L., Lip G.Y. Relationship between glucosamine use and the risk of lung cancer: Data from a nationwide prospective cohort study. Eur. Respir. J. 2022;59:2101399. doi: 10.1183/13993003.01399-2021. [DOI] [PubMed] [Google Scholar]
• 28.Quastel J.H., Cantero A. Inhibition of tumour growth by D-glucosamine. Nature. 1953;171:252–254. doi: 10.1038/171252a0. [DOI] [PubMed] [Google Scholar]
• 29.Molnar Z., Bekesi J.G. Cytotoxic effects of D-glucosamine on the ultrastructures of normal and neoplastic tissues in vivo. Cancer Res. 1972;32:756–765. [PubMed] [Google Scholar]
• 30.Friedman S.J., Skehan P. Membrane-active drugs potentiate the killing of tumor cells by D-glucosamine. Proc. Natl. Acad. Sci. USA. 1980;77:1172–1176. doi: 10.1073/pnas.77.2.1172. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 31.Oh H.-J., Lee J.S., Song D.-K., Shin D.-H., Jang B.-C., Suh S.-I., Park J.-W., Suh M.-H., Baek W.-K. D-glucosamine inhibits proliferation of human cancer cells through inhibition of p70S6K. Biochem. Biophys. Res. Commun. 2007;360:840–845. doi: 10.1016/j.bbrc.2007.06.137. [DOI] [PubMed] [Google Scholar]
• 32.Wang L.S., Chen S.J., Zhang J.F., Liu M.N., Zheng J.H., Yao X.D. Anti-proliferative potential of Glucosamine in renal cancer cells via inducing cell cycle arrest at G0/G1 phase. BMC Urol. 2017;17:38. doi: 10.1186/s12894-017-0221-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 33.de Keizer P.L., Packer L.M., Szypowska A.A., Riedl-Polderman P.E., van den Broek N.J., de Bruin A., Dansen T.B., Marais R., Brenkman A.B., Burgering B.M. Activation of forkhead box O transcription factors by oncogenic BRAF promotes p21cip1-dependent senescence. Cancer Res. 2010;70:8526–8536. doi: 10.1158/0008-5472.CAN-10-1563. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 34.Huang H., Tindall D.J. Dynamic FoxO transcription factors. Pt 15J. Cell Sci. 2007;120:2479–2487. doi: 10.1242/jcs.001222. [DOI] [PubMed] [Google Scholar]
• 35.Yu Z., Ju Y., Liu H. Anti-lung cancer effect of glucosamine by suppressing the phosphorylation of FOXO. Mol. Med. Rep. 2017;16:3395–3400. doi: 10.3892/mmr.2017.6976. [DOI] [PubMed] [Google Scholar]
• 36.Yu H., Jove R. The STATs of cancer--new molecular targets come of age. Nat. Rev. Cancer. 2004;4:97–105. doi: 10.1038/nrc1275. [DOI] [PubMed] [Google Scholar]
• 37.Chesnokov V., Sun C., Itakura K. Glucosamine suppresses proliferation of human prostate carcinoma DU145 cells through inhibition of STAT3 signaling. Cancer Cell Int. 2009;9:25. doi: 10.1186/1475-2867-9-25. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 38.Lin J.H., Walter P., Yen T.S. Endoplasmic reticulum stress in disease pathogenesis. Annu. Rev. Pathol. 2008;3:399–425. doi: 10.1146/annurev.pathmechdis.3.121806.151434. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 39.Morin M.J., Porter C.W., McKernan P., Bernacki R.J. The biochemical and ultrastructural effects of tunicamycin and D-glucosamine in L1210 leukemic cells. J. Cell Physiol. 1983;114:162–172. doi: 10.1002/jcp.1041140204. [DOI] [PubMed] [Google Scholar]
• 40.Werstuck G.H., Khan M.I., Femia G., Kim A.J., Tedesco V., Trigatti B., Shi Y. Glucosamine-induced endoplasmic reticulum dysfunction is associated with accelerated atherosclerosis in a hyperglycemic mouse model. Diabetes. 2006;55:93–101. doi: 10.2337/diabetes.55.01.06.db05-0633. [DOI] [PubMed] [Google Scholar]
• 41.Hwang M.-S., Baek W.-K. Glucosamine induces autophagic cell death through the stimulation of ER stress in human glioma cancer cells. Biochem. Biophys. Res. Commun. 2010;399:111–116. doi: 10.1016/j.bbrc.2010.07.050. [DOI] [PubMed] [Google Scholar]
• 42.Hientz K., Mohr A., Bhakta-Guha D., Efferth T. The role of p53 in cancer drug resistance and targeted chemotherapy. Oncotarget. 2017;8:8921–8946. doi: 10.18632/oncotarget.13475. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 43.Lefranc F., Facchini V., Kiss R. Proautophagic drugs: A novel means to combat apoptosis-resistant cancers, with a special emphasis on glioblastomas. Oncologist. 2007;12:1395–1403. doi: 10.1634/theoncologist.12-12-1395. [DOI] [PubMed] [Google Scholar]
• 44.Newens K., Walton J. A review of sugar consumption from nationally representative dietary surveys across the world. J. Hum. Nutr. Diet. 2016;29:225–240. doi: 10.1111/jhn.12338. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 45.Holesh J.E., Aslam S., Martin A. StatPearls. StatPearls Publishing; Treasure Island, FL, USA: 2023. [(accessed on 23 August 2023)]. Physiology, Carbohydrates. Available online: https://www.ncbi.nlm.nih.gov/books/NBK459280/ [Google Scholar]
• 46.Kilcoyne M., Joshi L. Carbohydrates in therapeutics. Cardiovasc. Hematol. Agents Med. Chem. 2007;5:186–197. doi: 10.2174/187152507781058663. [DOI] [PubMed] [Google Scholar]
• 47.Varki A., Sharon N. Essentials of Glycobiology. 2nd ed. Cold Spring Harbor Laboratory Press; Cold Spring Harbor, NY, USA: 2009. Historical Background and Overview. Chapter 1. [PubMed] [Google Scholar]
• 48.Iannetti E.F., Smeitink J.A.M., Willems P.H.G.M., Beyrath J., Koopman W.J.H. Rescue from galactose-induced death of Leigh Syndrome patient cells by pyruvate and NAD+ Cell Death Dis. 2018;9:1135. doi: 10.1038/s41419-018-1179-4. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 49.King M.W. Integrative Medical Biochemistry Examination and Board Review. McGraw-Hill Education; New York, NY, USA: 2014. Carbohydrates: Galactose Metabolism. [Google Scholar]
• 50.Conte F., van Buuringen N., Voermans N.C., Lefeber D.J. Galactose in human metabolism, glycosylation and congenital metabolic diseases: Time for a closer look. Biochim. Biophys. Acta Gen. Subj. 2021;1865:129898. doi: 10.1016/j.bbagen.2021.129898. [DOI] [PubMed] [Google Scholar]
• 51.Li N., He Y., Wang L., Mo C., Zhang J., Zhang W., Li J., Liao Z., Tang X., Xiao H. D-galactose induces necroptotic cell death in neuroblastoma cell lines. J. Cell Biochem. 2011;112:3834–3844. doi: 10.1002/jcb.23314. [DOI] [PubMed] [Google Scholar]
• 52.Shiratori R., Furuichi K., Yamaguchi M., Miyazaki N., Aoki H., Chibana H., Ito K., Aoki S. Glycolytic suppression dramatically changes the intracellular metabolic profile of multiple cancer cell lines in a mitochondrial metabolism-dependent manner. Sci. Rep. 2019;9:18699. doi: 10.1038/s41598-019-55296-3. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 53.Zheng D., Sussman J.H., Jeon M.P., Parrish S.T., MacMullan M.A., Delfarah A., Graham N.A. AKT but not MYC promotes reactive oxygen species-mediated cell death in oxidative culture. J. Cell Sci. 2020;133:jcs239277. doi: 10.1242/jcs.239277. [DOI] [PubMed] [Google Scholar]
• 54.Elstrom R.L., Bauer D.E., Buzzai M., Karnauskas R., Harris M.H., Plas D.R., Zhuang H., Cinalli R.M., Alavi A., Rudin C.M., et al. Akt Stimulates Aerobic Glycolysis in Cancer Cells. Cancer Res. 2004;64:3892–3899. doi: 10.1158/0008-5472.CAN-03-2904. [DOI] [PubMed] [Google Scholar]
• 55.Lanaspa M.A., Ishimoto T., Li N., Cicerchi C., Orlicky D.J., Ruzycki P., Rivard C., Inaba S., Roncal-Jimenez C.A., Bales E.S., et al. Endogenous fructose production and metabolism in the liver contributes to the development of metabolic syndrome. Nat. Commun. 2013;4:2434. doi: 10.1038/ncomms3434. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 56.Harris D.S., Slot J.W., Geuze H.J., James D.E. Polarized distribution of glucose transporter isoforms in Caco-2 cells. Proc. Natl. Acad. Sci. USA. 1992;89:7556–7560. doi: 10.1073/pnas.89.16.7556. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 57.Mahraoui L., Rousset M., Dussaulx E., Darmoul D., Zweibaum A., Brot-Laroche E., Jones H.F., Butler R.N., Brooks D.A., Nicola J.P., et al. Expression and localization of GLUT-5 in Caco-2 cells, human small intestine, and colon. Pt 1Am. J. Physiol. 1992;263:G312–G318. doi: 10.1152/ajpgi.1992.263.3.G312. [DOI] [PubMed] [Google Scholar]
• 58.Zamora-León S.P., Golde D.W., I Concha I., I Rivas C., Delgado-López F., Baselga J., Nualart F., Vera J.C. Expression of the fructose transporter GLUT5 in human breast cancer. Proc. Natl. Acad. Sci. USA. 1996;93:1847–1852. doi: 10.1073/pnas.93.5.1847. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 59.Nakagawa T., Lanaspa M.A., Millan I.S., Fini M., Rivard C.J., Sanchez-Lozada L.G., Andres-Hernando A., Tolan D.R., Johnson R.J. Fructose contributes to the Warburg effect for cancer growth. Cancer Metab. 2020;8:16. doi: 10.1186/s40170-020-00222-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 60.Nakagawa T., Tuttle K.R., A Short R., Johnson R.J. Hypothesis: Fructose-induced hyperuricemia as a causal mechanism for the epidemic of the metabolic syndrome. Nat. Clin. Pract. Nephrol. 2005;1:80–86. doi: 10.1038/ncpneph0019. [DOI] [PubMed] [Google Scholar]
• 61.Goncalves M.D., Lu C., Tutnauer J., Hartman T.E., Hwang S.-K., Murphy C.J., Pauli C., Morris R., Taylor S., Bosch K., et al. High-fructose corn syrup enhances intestinal tumor growth in mice. Science. 2019;363:1345–1349. doi: 10.1126/science.aat8515. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 62.Weng Y., Fan X., Bai Y., Wang S., Huang H., Yang H., Zhu J., Zhang F. SLC2A5 promotes lung adenocarcinoma cell growth and metastasis by enhancing fructose utilization. Cell Death Discov. 2018;4:38. doi: 10.1038/s41420-018-0038-5. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 63.Zwarts I., van Zutphen T., Kruit J.K., Liu W., Oosterveer M.H., Verkade H.J., Uhlenhaut N.H., Jonker J.W. Identification of the fructose transporter GLUT5 (SLC2A5) as a novel target of nuclear receptor LXR. Sci. Rep. 2019;9:9299. doi: 10.1038/s41598-019-45803-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 64.Godoy A., Ulloa V., Rodríguez F., Reinicke K., Yañez A.J., García M.D.L.A., Medina R.A., Carrasco M., Barberis S., Castro T., et al. Differential subcellular distribution of glucose transporters GLUT1–6 and GLUT9 in human cancer: Ultrastructural localization of GLUT1 and GLUT5 in breast tumor tissues. J. Cell Physiol. 2006;207:614–627. doi: 10.1002/jcp.20606. [DOI] [PubMed] [Google Scholar]
• 65.Karbassi M., Monzavi-Karbassi B., Hine R.J., Stanley J.S., Ramani V.P., Carcel-Trullols J., Whitehead T.L., Kelly T., Siegel E.R., Artaud C., et al. Fructose as a carbon source induces an aggressive phenotype in MDA-MB-468 breast tumor cells. Int. J. Oncol. 2010;37:615–622. doi: 10.3892/ijo_00000710. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 66.Wang X., Taniguchi N. Core Fucosylation of N-Linked Glycan for Fine-Tuning TGF b Receptor Function. Glycosci. Biol. Med. 2015;2014:991–997. [Google Scholar]
• 67.Zhao Y., Takahashi M., Gu J., Miyoshi E., Matsumoto A., Kitazume S., Taniguchi N. Functional roles of N-glycans in cell signaling and cell adhesion in cancer. Cancer Sci. 2008;99:1304–1310. doi: 10.1111/j.1349-7006.2008.00839.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 68.Listinsky J.J., Listinsky C.M., Alapati V., Siegal G.P. Cell surface fucose ablation as a therapeutic strategy for malignant neoplasms. Adv. Anat. Pathol. 2001;8:330–337. doi: 10.1097/00125480-200111000-00003. [DOI] [PubMed] [Google Scholar]
• 69.MacDougall S.L., Schwarting G.A., Parkinson D., Sullivan A.K. Increased fucosylation of glycolipids in a human leukaemia cell line (K562-Clone I) with decreased sensitivity to NK-mediated lysis. Immunology. 1987;62:75–80. [PMC free article] [PubMed] [Google Scholar]
• 70.Eccles M.R., Chatterjee A., Rodger E.J. Identifying drivers of metastasis; towards a systematic approach. Transl. Cancer Res. 2017;6:1273–1276. doi: 10.21037/tcr.2017.09.52. [DOI] [Google Scholar]
• 71.Vanhooren P.T., Vandamme E.J. L-Fucose: Occurrence, physiological role, chemical, enzymatic and microbial synthesis. J. Chem. Technol. Biotechnol. 1999;74:479–497. doi: 10.1002/(SICI)1097-4660(199906)74:6&#x0003c;479::AID-JCTB76&#x0003e;3.0.CO;2-E. [DOI] [Google Scholar]
• 72.Gorelik E., Xu F., Henion T., Anaraki F., Galili U. Reduction of metastatic properties of BL6 melanoma cells expressing terminal fucose(alpha)1-2-galactose after alpha1,2-fucosyltransferase cDNA transfection. Cancer Res. 1997;57:332–336. [PubMed] [Google Scholar]
• 73.Hotta H., Hamamura K., Yamashita K., Shibuya H., Tokuda N., Hashimoto N., Furukawa K., Yamamoto N., Hattori H., Toyokuni S., et al. Lewis y antigen is expressed in oral squamous cell carcinoma cell lines and tissues, but disappears in the invasive regions leading to the enhanced malignant properties irrespective of sialyl-Lewis x. Glycoconj J. 2013;30:585–597. doi: 10.1007/s10719-012-9458-2. [DOI] [PubMed] [Google Scholar]
• 74.Mathieu S., Prorok M., Benoliel A.-M., Uch R., Langlet C., Bongrand P., Gerolami R., El-Battari A. Transgene Expression of α(1,2)-Fucosyltransferase-I (FUT1) in Tumor Cells Selectively Inhibits Sialyl-Lewis x Expression and Binding to E-Selectin without Affecting Synthesis of Sialyl-Lewis a or Binding to P-Selectin. Am. J. Pathol. 2004;164:371–383. doi: 10.1016/S0002-9440(10)63127-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 75.Lester D.K., Burton C., Gardner A., Innamarato P., Kodumudi K., Liu Q., Adhikari E., Ming Q., Williamson D.B., Frederick D.T., et al. Fucosylation of HLA-DRB1 regulates CD4+ T cell-mediated anti-melanoma immunity and enhances immunotherapy efficacy. Nat. Cancer. 2023;4:222–239. doi: 10.1038/s43018-022-00506-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 76.Li Y., Jiang Y., Zhang L., Qian W., Hou X., Lin R. Exogenous l-fucose protects the intestinal mucosal barrier depending on upregulation of FUT2-mediated fucosylation of intestinal epithelial cells. Faseb J. 2021;35:e21699. doi: 10.1096/fj.202002446RRRR. [DOI] [PubMed] [Google Scholar]
• 77.Mullen J.L., Rosato F.E., Allen T.R., Miller E.E., Roseman J., Rosato E.F. Continuous intravenous fucose therapy in rat mammary cancer. II. J. Surg. Oncol. 1973;5:61–69. doi: 10.1002/jso.2930050109. [DOI] [PubMed] [Google Scholar]
• 78.Rosato F.E., Mullen J.L., Rosato E.F., Steiger E., Miller E. Continuous intravenous fucose treatment of rat mammary tumor. J. Surg. Oncol. 1972;4:94–101. doi: 10.1002/jso.2930040204. [DOI] [PubMed] [Google Scholar]
• 79.Roseman J.M., Miller E., Seltzer M.H., Wolfe D., Rosato F.E. The effect of L-fucose on rat mammary tumor growth. II. In vitro studies. J. Surg. Oncol. 1971;3:79–88. doi: 10.1002/jso.2930030114. [DOI] [PubMed] [Google Scholar]
• 80.Tomsik P., Soukup T., Cermakova E., Micuda S., Niang M., Sucha L., Rezacova M. L-rhamnose and L-fucose suppress cancer growth in mice. Cent. Eur. J. Biol. 2011;6:1–9. doi: 10.2478/s11535-010-0087-0. [DOI] [Google Scholar]
• 81.Yao Y., Qian C., Chang Z., Yang J., Chen Y., Wang H., Zhu J., Xiao Y., Li Y., Zhao J., et al. L-Fucose increases the fucosylation of colorectal cancer cells via promoting the accumulation of serine. Food Funct. 2023;14:4314–4326. doi: 10.1039/D3FO00820G. [DOI] [PubMed] [Google Scholar]
• 82.Christiansen M.N., Chik J., Lee L., Anugraham M., Abrahams J.L., Packer N.H. Cell surface protein glycosylation in cancer. Proteomics. 2014;14:525–546. doi: 10.1002/pmic.201300387. [DOI] [PubMed] [Google Scholar]
• 83.Kudelka M.R., Stowell S.R., Cummings R.D., Neish A.S. Intestinal epithelial glycosylation in homeostasis and gut microbiota interactions in IBD. Nat. Rev. Gastroenterol. Hepatol. 2020;17:597–617. doi: 10.1038/s41575-020-0331-7. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 84.Izumori K. Bioproduction strategies for rare hexose sugars. Sci. Nat. 2002;89:120–124. doi: 10.1007/s00114-002-0297-z. [DOI] [PubMed] [Google Scholar]
• 85.Smith A., Avery A., Ford R., Yang Q., Goux A., Mukherjee I., Neville D.C.A., Jethwa P.H. Rare sugars: Metabolic impacts and mechanisms of action: A scoping review. Br. J. Nutr. 2021;128:389–406. doi: 10.1017/S0007114521003524. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 86.Umai D., Kayalvizhi R., Kumar V., Jacob S. Xylitol: Bioproduction and Applications—A Review. Front. Sustain. 2022;3:826190. doi: 10.3389/frsus.2022.826190. [DOI] [Google Scholar]
• 87.Chukwuma C.I., Islam M.S. Effects of xylitol on carbohydrate digesting enzymes activity, intestinal glucose absorption and muscle glucose uptake: A multi-mode study. Food Funct. 2015;6:955–962. doi: 10.1039/C4FO00994K. [DOI] [PubMed] [Google Scholar]
• 88.Chukwuma C., Islam M. Sweetners: Reference Series in Phytochemistry. Springer; New York, NY, USA: 2016. Xylitol: One Name, Numerous Benefits; pp. 1–27. [Google Scholar]
• 89.Miyasawa-Hori H., Aizawa S., Takahashi N. Difference in the xylitol sensitivity of acid production among Streptococcus mutans strains and the biochemical mechanism. Oral Microbiol. Immunol. 2006;21:201–205. doi: 10.1111/j.1399-302X.2006.00273.x. [DOI] [PubMed] [Google Scholar]
• 90.Sato J., Wang Y.M., van Eys J. Metabolism of xylitol and glucose in rats bearing hepatocellular. Cancer Res. 1981;41:3192–3199. [PubMed] [Google Scholar]
• 91.Ahuja V., Macho M., Ewe D., Singh M., Saha S., Saurav K. Biological and Pharmacological Potential of Xylitol: A Molecular Insight of Unique Metabolism. Foods. 2020;9:1592. doi: 10.3390/foods9111592. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 92.Park E., Park M.H., Na H.S., Chung J. Xylitol induces cell death in lung cancer A549 cells by autophagy. Biotechnol. Lett. 2015;37:983–990. doi: 10.1007/s10529-014-1757-1. [DOI] [PubMed] [Google Scholar]
• 93.Trachootham D., Chingsuwanrote P., Yoosadiang P., Mekkriangkrai D., Ratchawong T., Buraphacheep N., Kijanukul S., Saekhow S., Pongpitchayadej O., Vongvachvasin K., et al. Partial Substitution of Glucose with Xylitol Suppressed the Glycolysis and Selectively Inhibited the Proliferation of Oral Cancer Cells. Nutr. Cancer. 2017;69:862–872. doi: 10.1080/01635581.2017.1339097. [DOI] [PubMed] [Google Scholar]
• 94.Sahasakul Y., Angkhasirisap W., Lam-Ubol A., Aursalung A., Sano D., Takada K., Trachootham D. Partial Substitution of Glucose with Xylitol Prolongs Survival and Suppresses Cell Proliferation and Glycolysis of Mice Bearing Orthotopic Xenograft of Oral Cancer. Nutrients. 2022;14:2023. doi: 10.3390/nu14102023. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 95.Tomonobu N., Komalasari N.L.G.Y., Sumardika I.W., Jiang F., Chen Y., Yamamoto K.I., Kinoshita R., Murata H., Inoue Y., Sakaguchi M. Xylitol acts as an anticancer monosaccharide to induce selective cancer death via regulation of the glutathione level. Chem. Biol. Interact. 2020;324:109085. doi: 10.1016/j.cbi.2020.109085. [DOI] [PubMed] [Google Scholar]
• 96.Liu Y., Wu D., Fu Q., Hao S., Gu Y., Zhao W., Chen S., Sheng F., Xu Y., Chen Z., et al. CHAC1 as a Novel Contributor of Ferroptosis in Retinal Pigment Epithelial Cells with Oxidative Damage. Int. J. Mol. Sci. 2023;24:1582. doi: 10.3390/ijms24021582. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 97.Iga Y., Nakamichi K., Shirai Y., Matsuo T. Acute and sub-chronic toxicity of D-allose in rats. Biosci. Biotechnol. Biochem. 2010;74:1476–1478. doi: 10.1271/bbb.100121. [DOI] [PubMed] [Google Scholar]
• 98.Weckwerth W., Loureiro M.E., Wenzel K., Fiehn O. Differential metabolic networks unravel the effects of silent plant phenotypes. Proc. Natl. Acad. Sci. USA. 2004;101:7809–7814. doi: 10.1073/pnas.0303415101. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 99.Kannan R.R.R., Arumugam R., Anantharaman P. Anantharaman, Chemical composition and antibacterial activity of Indian seagrasses against urinary tract pathogens. Food Chem. 2012;135:2470–2473. doi: 10.1016/j.foodchem.2012.07.070. [DOI] [PubMed] [Google Scholar]
• 100.Chen Z., Chen J., Zhang W., Zhang T., Guang C., Mu W. Recent research on the physiological functions, applications, and biotechnological production of d-allose. Appl. Microbiol. Biotechnol. 2018;102:4269–4278. doi: 10.1007/s00253-018-8916-6. [DOI] [PubMed] [Google Scholar]
• 101.Sui L., Dong Y., Watanabe Y., Yamaguchi F., Hatano N., Tsukamoto I., Izumori K., Tokuda M. The inhibitory effect and possible mechanisms of D-allose on cancer cell proliferation. Int. J. Oncol. 2005;27:907–912. doi: 10.3892/ijo.27.4.907. [DOI] [PubMed] [Google Scholar]
• 102.Sui L., Dong Y., Watanabe Y., Yamaguchi F., Hatano N., Izumori K., Tokuda M. Growth inhibitory effect of D-allose on human ovarian carcinoma cells in vitro. Anticancer Res. 2005;25:2639–2644. [PubMed] [Google Scholar]
• 103.Jeong R.U., Lim S., Kim M.O., Moon M.H. Effect of D-allose on prostate cancer cell lines: Phospholipid profiling by nanoflow liquid chromatography-tandem mass spectrometry. Anal. Bioanal. Chem. 2011;401:689–698. doi: 10.1007/s00216-011-5113-1. [DOI] [PubMed] [Google Scholar]
• 104.Hirata Y., Saito M., Tsukamoto I., Yamaguchi F., Sui L., Kamitori K., Dong Y., Uehara E., Konishi R., Janjua N., et al. Analysis of the inhibitory mechanism of D-allose on MOLT-4F leukemia cell proliferation. J. Biosci. Bioeng. 2009;107:562–568. doi: 10.1016/j.jbiosc.2008.12.021. [DOI] [PubMed] [Google Scholar]
• 105.Han S.H., Jeon J.H., Ju H.R., Jung U., Kim K.Y., Yoo H.S., Lee Y.H., Song K.S., Hwang H.M., Na Y.S., et al. VDUP1 upregulated by TGF-β1 and 1,25-dihydorxyvitamin D3 inhibits tumor cell growth by blocking cell-cycle progression. Oncogene. 2003;22:4035–4046. doi: 10.1038/sj.onc.1206610. [DOI] [PubMed] [Google Scholar]
• 106.Ohta S., Lai E.W., Pang A.L., Brouwers F.M., Chan W.-Y., Eisenhofer G., de Krijger R., Ksinantova L., Breza J., Blazicek P., et al. Downregulation of metastasis suppressor genes in malignant pheochromocytoma. Int. J. Cancer. 2005;114:139–143. doi: 10.1002/ijc.20670. [DOI] [PubMed] [Google Scholar]
• 107.Yamaguchi F., Takata M., Kamitori K., Nonaka M., Dong Y., Sui L., Tokuda M. Rare sugar D-allose induces specific up-regulation of TXNIP and subsequent G1 cell cycle arrest in hepatocellular carcinoma cells by stabilization of p27kip1. Int. J. Oncol. 2008;32:377–385. doi: 10.3892/ijo.32.2.377. [DOI] [PubMed] [Google Scholar]
• 108.Parikh H., Carlsson E., A Chutkow W., E Johansson L., Storgaard H., Poulsen P., Saxena R., Ladd C., Schulze P.C., Mazzini M.J., et al. TXNIP regulates peripheral glucose metabolism in humans. PLoS Med. 2007;4:e158. doi: 10.1371/journal.pmed.0040158. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 109.Hoshikawa H., Mori T., Mori N. In vitro and in vivo effects of D-allose: Up-regulation of thioredoxin-interacting protein in head and neck cancer cells. Ann. Otol. Rhinol. Laryngol. 2010;119:567–571. doi: 10.1177/000348941011900810. [DOI] [PubMed] [Google Scholar]
• 110.Noguchi C., Kamitori K., Hossain A., Hoshikawa H., Katagi A., Dong Y., Sui L., Tokuda M., Yamaguchi F. D-Allose Inhibits Cancer Cell Growth by Reducing GLUT1 Expression. Tohoku J. Exp. Med. 2016;238:131–141. doi: 10.1620/tjem.238.131. [DOI] [PubMed] [Google Scholar]
• 111.Naha N., Lee H.Y., Jo M.J., Chung B.C., Kim S.H., Kim M.O. Rare sugar D-allose induces programmed cell death in hormone refractory prostate cancer cells. Apoptosis. 2008;13:1121–1134. doi: 10.1007/s10495-008-0232-7. [DOI] [PubMed] [Google Scholar]
• 112.Torrealba N., Rodríguez-Berriguete G., Vera R., Fraile B., Olmedilla G., Martínez-Onsurbe P., Sánchez-Chapado M., Paniagua R., Royuela M. Homeostasis: Apoptosis and cell cycle in normal and pathological prostate. Aging Male. 2020;23:335–345. doi: 10.1080/13685538.2018.1470233. [DOI] [PubMed] [Google Scholar]
• 113.Kishida K., Iida T., Yamada T., Toyoda Y. Intestinal absorption of D-fructose isomers, D-allulose, D-sorbose and D-tagatose, via glucose transporter type 5 (GLUT5) but not sodium-dependent glucose cotransporter 1 (SGLT1) in rats. Br. J. Nutr. 2023;130:1852–1858. doi: 10.1017/S0007114523001113. [DOI] [PubMed] [Google Scholar]
• 114.Zebiri I., Balieu S., Guilleret A., Reynaud R., Haudrechy A. The Chemistry of L-Sorbose. Eur. J. Org. Chem. 2011;2011:2905–2910. doi: 10.1002/ejoc.201001578. [DOI] [Google Scholar]
• 115.Moore D., Stewart G.R. Effects of 2-Deoxy-d-Glucose, d-Glucosamine, and l-Sorbose on the Growth of Coprinus lagopus hyphae. J. Gen. Microbiol. 1972;71:333–342. doi: 10.1099/00221287-71-2-333. [DOI] [Google Scholar]
• 116.Sahoo D.K., Mishra S., Bisaria V.S. Influence of L-Sorbose on Growth and Enzyme Synthesis of Trichoderma reesei C-5. Microbiology. 1986;132:2761–2766. doi: 10.1099/00221287-132-10-2761. [DOI] [Google Scholar]
• 117.Noronha J.C., Braunstein C.R., Mejia S.B., Khan T.A., Kendall C.W.C., Wolever T.M.S., Leiter L.A., Sievenpiper J.L. The Effect of Small Doses of Fructose and Its Epimers on Glycemic Control: A Systematic Review and Meta-Analysis of Controlled Feeding Trials. Nutrients. 2018;10:1805. doi: 10.3390/nu10111805. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 118.Xu H.-L., Zhou X., Chen S., Xu S., Li Z., Nakanishi H., Gao X.-D. Rare sugar l-sorbose exerts antitumor activity by impairing glucose metabolism. Commun. Biol. 2023;6:259. doi: 10.1038/s42003-023-04638-z. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 119.Xu D., Li X., Shao F., Lv G., Lv H., Lee J.-H., Qian X., Wang Z., Xia Y., Du L., et al. The protein kinase activity of fructokinase A specifies the antioxidant responses of tumor cells by phosphorylating p62. Sci. Adv. 2019;5:eaav4570. doi: 10.1126/sciadv.aav4570. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 120.Giraud M.-F., Naismith J.H. The rhamnose pathway. Curr. Opin. Struct. Biol. 2000;10:687–696. doi: 10.1016/S0959-440X(00)00145-7. [DOI] [PubMed] [Google Scholar]
• 121.Sheridan R.T.C., Hudon J., Hank J.A., Sondel P.M., Kiessling L.L. Rhamnose glycoconjugates for the recruitment of endogenous anti-carbohydrate antibodies to tumor cells. Chembiochem. 2014;15:1393–1398. doi: 10.1002/cbic.201402019. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 122.Tomšík P., Stoklasová A., Mičuda S., Niang M., Šuba P., Knížek J., Řezáčová M. Evaluation of the Antineoplastic Activity of L-rhamnose in vitro. A Comparison with 2-deoxyglucose. Acta Med. 2008;51:113–119. doi: 10.14712/18059694.2017.12. [DOI] [PubMed] [Google Scholar]
• 123.Li X., Rao X., Cai L., Liu X., Wang H., Wu W., Zhu C., Chen M., Wang P.G., Yi W. Targeting Tumor Cells by Natural Anti-Carbohydrate Antibodies Using Rhamnose-Functionalized Liposomes. ACS Chem. Biol. 2016;11:1205–1209. doi: 10.1021/acschembio.6b00173. [DOI] [PubMed] [Google Scholar]
• 124.Li S., Chen F., Li Y., Wang L., Li H., Gu G., Li E. Rhamnose-Containing Compounds: Biosynthesis and Applications. Molecules. 2022;27:5315. doi: 10.3390/molecules27165315. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 125.Wijayasinghe Y.S., Bhansali M.P., Borkar M.R., Chaturbhuj G.U., Muntean B.S., Viola R.E., Bhansali P.R. A Comprehensive Biological and Synthetic Perspective on 2-Deoxy-d-Glucose (2-DG), A Sweet Molecule with Therapeutic and Diagnostic Potentials. J. Med. Chem. 2022;65:3706–3728. doi: 10.1021/acs.jmedchem.1c01737. [DOI] [PubMed] [Google Scholar]
• 126.Singh R., Gupta V., Kumar A., Singh K. 2-Deoxy-D-Glucose: A Novel Pharmacological Agent for Killing Hypoxic Tumor Cells, Oxygen Dependence-Lowering in COVID-19, and Other Pharmacological Activities. Adv. Pharmacol. Pharm. Sci. 2023;2023:9993386. doi: 10.1155/2023/9993386. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 127.Laussel C., Léon S. Cellular toxicity of the metabolic inhibitor 2-deoxyglucose and associated resistance mechanisms. Biochem. Pharmacol. 2020;182:114213. doi: 10.1016/j.bcp.2020.114213. [DOI] [PubMed] [Google Scholar]
• 128.Wick A.N., Drury D.R., Nakada H.I., Wolfe J.B., Britton B., Grabowski R. Localization of the primary metabolic block produced by 2-deoxyglucose. J. Biol. Chem. 1957;224:963–969. doi: 10.1016/S0021-9258(18)64988-9. [DOI] [PubMed] [Google Scholar]
• 129.Chen W., Gueron M.J.B. The inhibition of bovine heart hexokinase by 2-deoxy-D-glucose-6-phosphate: Characterization by 31P NMR and metabolic implications. Biochimie. 1992;74:867–873. doi: 10.1016/0300-9084(92)90070-U. [DOI] [PubMed] [Google Scholar]
• 130.Liu H., Savaraj N., Priebe W., Lampidis T.J. Hypoxia increases tumor cell sensitivity to glycolytic inhibitors: A strategy for solid tumor therapy (Model C) Biochem. Pharmacol. 2002;64:1745–1751. doi: 10.1016/S0006-2952(02)01456-9. [DOI] [PubMed] [Google Scholar]
• 131.Yun C.W., Lee S.H. The Roles of Autophagy in Cancer. Int. J. Mol. Sci. 2018;19:3466. doi: 10.3390/ijms19113466. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 132.Kurtoglu M., Gao N., Shang J., Maher J.C., Lehrman M.A., Wangpaichitr M., Savaraj N., Lane A.N., Lampidis T.J. Under normoxia, 2-deoxy-D-glucose elicits cell death in select tumor types not by inhibition of glycolysis but by interfering with N-linked glycosylation. Mol. Cancer Ther. 2007;6:3049–3058. doi: 10.1158/1535-7163.MCT-07-0310. [DOI] [PubMed] [Google Scholar]
• 133.Datema R., Schwarz R.T., Jankowski A.W. Fluoroglucose-inhibition of protein glycosylation in vivo. Inhibition of mannose and glucose incorporation into lipid-linked oligosaccharides. Eur. J. Biochem. 1980;109:331–341. doi: 10.1111/j.1432-1033.1980.tb04799.x. [DOI] [PubMed] [Google Scholar]
• 134.Datema R., Schwarz R.T. Interference with glycosylation of glycoproteins. Inhibition of formation of lipid-linked oligosaccharides in vivo. Biochem. J. 1979;184:113–123. doi: 10.1042/bj1840113. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 135.Schmidt M.F.G., Biely P., Kratky Z., Schwarz R.T. Metabolism of 2-deoxy-2-fluoro-D-[3H]glucose and 2-deoxy-2-fluoro-D-[3H]mannose in yeast and chick-embryo cells. Eur. J. Biochem. 1978;87:55–68. doi: 10.1111/j.1432-1033.1978.tb12351.x. [DOI] [PubMed] [Google Scholar]
• 136.Schmidt M.F., Schwarz R.T., Scholtissek C. Nucleoside-diphosphate derivatives of 2-deoxy-D-glucose in animal cells. Eur. J. Biochem. 1974;49:237–247. doi: 10.1111/j.1432-1033.1974.tb03828.x. [DOI] [PubMed] [Google Scholar]
• 137.Ramírez-Peinado S., Alcázar-Limones F., Lagares-Tena L., El Mjiyad N., Caro-Maldonado A., Tirado O.M., Muñoz-Pinedo C. 2-deoxyglucose induces Noxa-dependent apoptosis in alveolar rhabdomyosarcoma. Cancer Res. 2011;71:6796–6806. doi: 10.1158/0008-5472.CAN-11-0759. [DOI] [PubMed] [Google Scholar]
• 138.Stein M., Lin H., Jeyamohan C., Dvorzhinski D., Gounder M., Bray K., Eddy S., Goodin S., White E., Dipaola R.S. Targeting tumor metabolism with 2-deoxyglucose in patients with castrate-resistant prostate cancer and advanced malignancies. Prostate. 2010;70:1388–1394. doi: 10.1002/pros.21172. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 139.Wu H., Zhu H., Liu D.X., Niu T.-K., Ren X., Patel R., Hait W.N., Yang J.-M. Silencing of elongation factor-2 kinase potentiates the effect of 2-deoxy-D-glucose against human glioma cells through blunting of autophagy. Cancer Res. 2009;69:2453–2460. doi: 10.1158/0008-5472.CAN-08-2872. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 140.Xi H., Barredo J.C., Merchan J.R., Lampidis T.J. Endoplasmic reticulum stress induced by 2-deoxyglucose but not glucose starvation activates AMPK through CaMKKβ leading to autophagy. Biochem. Pharmacol. 2013;85:1463–1477. doi: 10.1016/j.bcp.2013.02.037. [DOI] [PubMed] [Google Scholar]
• 141.Pradelli L.A., Bénéteau M., Chauvin C., A Jacquin M., Marchetti S., Muñoz-Pinedo C., Auberger P., Pende M., Ricci J.-E. Glycolysis inhibition sensitizes tumor cells to death receptors-induced apoptosis by AMP kinase activation leading to Mcl-1 block in translation. Oncogene. 2010;29:1641–1652. doi: 10.1038/onc.2009.448. [DOI] [PubMed] [Google Scholar]
• 142.Zou Z., Tao T., Li H., Zhu X. mTOR signaling pathway and mTOR inhibitors in cancer: Progress and challenges. Cell Biosci. 2020;10:31. doi: 10.1186/s13578-020-00396-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 143.Tu D., Gao Y., Yang R., Guan T., Hong J.-S., Gao H.-M. The pentose phosphate pathway regulates chronic neuroinflammation and dopaminergic neurodegeneration. J. Neuroinflamm. 2019;16:255. doi: 10.1186/s12974-019-1659-1. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 144.Coleman M.C., Asbury C.R., Daniels D., Du J., Aykin-Burns N., Smith B.J., Li L., Spitz D.R., Cullen J.J. 2-deoxy-D-glucose causes cytotoxicity, oxidative stress, and radiosensitization in pancreatic cancer. Free Radic. Biol. Med. 2008;44:322–331. doi: 10.1016/j.freeradbiomed.2007.08.032. [DOI] [PubMed] [Google Scholar]
• 145.Xi H., Kurtoglu M., Lampidis T.J. The wonders of 2-deoxy-d-glucose. IUBMB Life. 2014;66:110–121. doi: 10.1002/iub.1251. [DOI] [PubMed] [Google Scholar]
• 146.Landau B.R., Laszlo J., Stengle J., Burk D. Certain metabolic and pharmacologic effects in cancer patients given infusions of 2-deoxy-D-glucose. J. Natl. Cancer Inst. 1958;21:485–494. [PubMed] [Google Scholar]
• 147.Fowler J.S., Ido T. Initial and subsequent approach for the synthesis of 18FDG. Nucl. Med. Semin. 2002;32:6–12. doi: 10.1053/snuc.2002.29270. [DOI] [PubMed] [Google Scholar]
• 148.Lampidis T.J., Kurtoglu M., Maher J.C., Liu H., Krishan A., Sheft V., Szymanski S., Fokt I., Rudnicki W.R., Ginalski K., et al. Efficacy of 2-halogen substituted d-glucose analogs in blocking glycolysis and killing “hypoxic tumor cells”. Cancer Chemother. Pharmacol. 2006;58:725–734. doi: 10.1007/s00280-006-0207-8. [DOI] [PubMed] [Google Scholar]
• 149.Priebe W., Zielinski R., Fokt I., Felix E., Radjendirane V., Arumugam J., Khuong M.T., Krasinski M., Skora S. EXTH-07. Design and Evaluation of wp1122, an Inhibitor of Glycolysis with Increased Cns Uptake. Neuro-Oncol. 2018;20((Suppl. S6)):vi86. doi: 10.1093/neuonc/noy148.356. [DOI] [Google Scholar]
• 150.Mokhtari R.B., Homayouni T.S., Baluch N., Morgatskaya E., Kumar S., Das B., Yeger H. Combination therapy in combating cancer. Oncotarget. 2017;8:38022–38043. doi: 10.18632/oncotarget.16723. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 151.Kim D.-S., Park K.-S., Jeong K.-C., Lee B.I., Lee C.-H., Kim S.-Y. Glucosamine is an effective chemo-sensitizer via transglutaminase 2 inhibition. Cancer Lett. 2009;273:243–249. doi: 10.1016/j.canlet.2008.08.015. [DOI] [PubMed] [Google Scholar]
• 152.Indo K., Hoshikawa H., Kamitori K., Yamaguchi F., Mori T., Tokuda M., Mori N. Effects of D-allose in combination with docetaxel in human head and neck cancer cells. Int. J. Oncol. 2014;45:2044–2050. doi: 10.3892/ijo.2014.2590. [DOI] [PubMed] [Google Scholar]
• 153.Shenfield G.M. Fixed Combination Drug Therapy. Drugs. 1982;23:462–480. doi: 10.2165/00003495-198223060-00003. [DOI] [PubMed] [Google Scholar]
• 154.Ekholm F.S., Berényi Á., Lagerquist L., Saloranta T., Zupkó I., Schneider G., Wölfling J., Leino R. Cytotoxic activity of some glycoconjugates including saponins and anthracyclines. Carbohydr. Res. 2012;356:295–298. doi: 10.1016/j.carres.2012.03.017. [DOI] [PubMed] [Google Scholar]
• 155.La Ferla B., Airoldi C., Zona C., Orsato A., Cardona F., Merlo S., Sironi E., D’Orazio G., Nicotra F. Natural glycoconjugates with antitumor activity. Nat. Prod. Rep. 2011;28:630–648. doi: 10.1039/C0NP00055H. [DOI] [PubMed] [Google Scholar]
• 156.Govindarajan M. Amphiphilic glycoconjugates as potential anti-cancer chemotherapeutics. Eur. J. Med. Chem. 2018;143:1208–1253. doi: 10.1016/j.ejmech.2017.10.015. [DOI] [PubMed] [Google Scholar]
• 157.Molejon M.I., Weiz G., Breccia J.D., Vaccaro M.I. Glycoconjugation: An approach to cancer therapeutics. World J. Clin. Oncol. 2020;11:110–120. doi: 10.5306/wjco.v11.i3.110. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 158.Pastuch-Gawołek G., Szreder J., Domińska M., Pielok M., Cichy P., Grymel M. A Small Sugar Molecule with Huge Potential in Targeted Cancer Therapy. Pharmaceutics. 2023;15:913. doi: 10.3390/pharmaceutics15030913. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 159.Halmos T., Santarromana M., Antonakis K., Scherman D. Synthesis of glucose-chlorambucil derivatives and their recognition by the human GLUT1 glucose transporter. Eur. J. Pharmacol. 1996;318:477–484. doi: 10.1016/S0014-2999(96)00796-0. [DOI] [PubMed] [Google Scholar]
• 160.Kumar P., Shustov G., Liang H., Khlebnikov V., Zheng W., Yang X.H., Cheeseman C., Wiebe L.I. Design, Synthesis, and Preliminary Biological Evaluation of 6-O-Glucose–Azomycin Adducts for Diagnosis and Therapy of Hypoxic Tumors. J. Med. Chem. 2012;55:6033–6046. doi: 10.1021/jm2017336. [DOI] [PubMed] [Google Scholar]
• 161.Ma Y., Chen H., Su S., Wang T., Zhang C., Fida G., Cui S., Zhao J., Gu Y. Galactose as Broad Ligand for Multiple Tumor Imaging and Therapy. J. Cancer. 2015;6:658–670. doi: 10.7150/jca.11647. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 162.Valieva Y., Ivanova E., Fayzullin A., Kurkov A., Igrunkova A. Senescence-Associated β-Galactosidase Detection in Pathology. Diagnostics. 2022;12:2309. doi: 10.3390/diagnostics12102309. [DOI] [PMC free article] [PubMed] [Google Scholar]
• 163.Meng X., Lian X., Li X., Ya Q., Li T., Zhang Y., Yang Y., Zhang Y. Synthesis of 2′-paclitaxel 2-deoxy-2-fluoro-glucopyranosyl carbonate for specific targeted delivery to cancer cells. Carbohydr. Res. 2020;493:108034. doi: 10.1016/j.carres.2020.108034. [DOI] [PubMed] [Google Scholar]
• 164.Najlah M. Drug repurposing supported by nanotechnology: A promising strategy to fight cancer. Ther. Deliv. 2021;12:267–269. doi: 10.4155/tde-2021-0009. [DOI] [PubMed] [Google Scholar]