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. 2025 Aug 7;15:28852. doi: 10.1038/s41598-025-13635-7

Giardia duodenalis stabilizes HIF-1α and induces glycolytic alterations in intestinal epithelial cells

Emily DeMichele 1,2,3,4, Olivia Sosnowski 1,2,3, Darragh Flood 4, Cormac T Taylor 4, Ian A Lewis 1, Thibault Allain 1,2,3,5,✉, Andre G Buret 1,2,3
PMCID: PMC12328747  PMID: 40770382

Abstract

The gastrointestinal epithelium relies on activation of the hypoxia-inducible factor (HIF) to promote cell survival and maintain bioenergetic homeostasis during hypoxia. While many pathogens can activate HIF, the effects of enteric protozoa on HIF activation in gastrointestinal epithelial cells remain unclear. Giardia duodenalis, a prevalent protozoan enteropathogen, causes intestinal barrier dysfunction characterized by epithelial malabsorption, mucus depletion, altered mucin glycosylation, and microbiota dysbiosis. Findings from the present study reveal an epithelial hypoxic signature upon Giardia infection. Human intestinal epithelial cells were exposed to vehicle or Giardia duodenalis isolate GS/M under normoxic (21% O2) or hypoxic (1% O2) conditions. In normoxia, infected cells displayed a time-dependent increase in HIF-1α protein expression, the oxygen-dependent subunit of HIF-1. In normoxia, Giardia infection upregulated HIF-1 target genes involved in cellular stress (i.e., VEGFA, ANKRD37, GADD45A) and glycolysis (i.e., HK2, LDHA). This was accompanied by changes in the abundance of glycolytic intermediates (i.e., glucose-6-phosphate, pyruvate, lactate). Although infection in hypoxia failed to augment the hypoxia-induced HIF-1α stabilization, HIF-1 target genes were still upregulated, albeit to a lesser degree. These findings indicate that Giardia induces a transient epithelial hypoxic response in normoxic conditions, revealing a hitherto unrecognized epithelial rescue response to this intestinal parasite.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-025-13635-7.

Keywords: Giardia, Giardiasis, Intestinal epithelial cells, Hypoxia, HIF-1α, Glycolysis

Subject terms: Parasite host response, Parasitology, Infection, Microbiology

Introduction

The protozoan parasite Giardia duodenalis (syn G. lamblia, G. intestinalis) is the causative agent of giardiasis, one of the most prevalent foodborne or waterborne diarrheal diseases worldwide. The prevalence of giardiasis is highest in developing countries, where it is estimated that nearly one-third of the population will become infected with the parasite during their lifetime1. Giardia is transmitted via the fecal-oral route, and ingesting 10 infective cysts is sufficient to cause an infection in mammalian hosts2,3. Once the metabolically dormant cysts reach the duodenum, they undergo excystation and release the active trophozoites that can interact with and disturb the resident microbiota, disrupt mucus and mucin glycosylation, and damage small intestinal epithelial function and integrity4–13. Importantly, the trophozoites are microaerophilic and are therefore well-suited to the small intestinal lumen where oxygen concentrations may fluctuate but remain between 1 and 3% O214–16. Giardia infections can remain asymptomatic or may cause clinical symptoms including diarrhea, steatorrhea, abdominal pain, flatulence, malabsorption, nausea, and/or vomiting4,17–19. Giardiasis can also lead to a variety of post-infectious complications, including post-infectious irritable bowel syndrome and extraintestinal pathologies20. Of note, only two of the eight genetic G. duodenalis Assembalges, A and B, are zoonotic and have been isolated from human infections21. While lacking epidemiological data has made it difficult to understand the differences in virulence between Assemblages A and B, the higher global prevalence of Assemblage B may be attributed to the higher parasitic load of these isolates22.

The important role of oxygen homeostasis in the gastrointestinal tract has become clear in the past decade, especially in the context of infection and inflammation. During an infection, tissues become hypoxic due to a combination of reduced oxygen availability, matched with increased oxygen consumption by infiltrating neutrophils and other immune cells23. To combat this environmental stressor, mammalian cells utilize the hypoxic inducible factor (HIF) to activate a transcriptional response that promotes cellular survival during hypoxia24. HIF consists of two constitutively translated subunits, but the dimerization of these subunits is controlled in an oxygen-dependent manner. The alpha subunit resides in the cytoplasm wherein it is hydroxylated by prolyl hydroxylases (PHDs) and Factor Inhibiting HIF (FIH) when cytoplasmic oxygen is abundant, leading to the proteasomal degradation and inactivation of HIF-1α25,26. In contrast, the lack of cytoplasmic oxygen during hypoxia enables the alpha subunit to evade hydroxylation and translocate to the nucleus where it will dimerize with the beta subunit and bind associated cofactors27. The HIF complex will recognize and bind to the hypoxia response element (HRE) sequence found in the promoter of hundreds of genes, driving a transcriptional response that modulates cellular processes such as erythropoiesis, angiogenesis, and cellular proliferation28. In addition, the hypoxic response helps maintain bioenergetic homeostasis by compensating for the loss in oxidative phosphorylation capacity. These compensatory mechanisms include HIF-1-induced changes in glycolytic gene expression29.

Although little is known regarding the role of HIF-1 during enteric protozoan infections, growing evidence suggests HIF-1 may have a role in cellular responses to Giardia infection30. Transcriptomic analyses of human organoid monolayers infected with G. duodenalis isolate WB (Assemblage A) revealed an enrichment of hypoxia-response-related genes up to 24 h after infection31. Similarly, transcriptomic analyses of human Caco-2 intestinal epithelial cells infected with G. duodenalis isolate WB or GS/M (Assemblage B) found that HIF-1 target genes were activated (i.e., NOS2, ANKRD37, GADD45A, ITF, MIR210HG, SLC2A3), consistent with the recent observation that G. duodenalis isolate WB extracellular vesicles could increase expression of HIF-1-associated genes in these cells32–34. These transcriptional signatures indicate a hypoxic cellular response is likely at play during infection, but the role of HIF-1 remains unknown. In the present study, we investigated the role of HIF-1α during G. duodenalis infection with an Assemblage B isolate (GS/M) to better characterize the epithelial response to enteropathogens. We hypothesized that Giardia induces HIF-1α stabilization in intestinal epithelial cells, thereby upregulating genes related to cellular stress and central carbon metabolism. Our results describe a novel host phenotype wherein Giardia drives a transient hypoxic response in normoxic intestinal epithelial cells, characterized by increased HIF-1α protein expression, as well as increased transcription of HIF-1 target genes associated with cell stress and glycolytic metabolism.

Results

Giardia stabilizes HIF-1α in normoxic intestinal epithelial cells

Non-differentiated Caco-2 intestinal epithelial cells were incubated for 1.5–4.5 h in normoxic (21% O2) or hypoxic (1% O2) conditions to assess the early and peak stabilization of HIF-1α in the context of Giardia infection. To validate the stabilization of HIF-1α by hypoxia, both the Stemcell hypoxia chamber and O2 control glove box systems were used. Both systems showed an early and peak protein expression of HIF-1α at 1.5 and 4.5 h, respectively (Fig. S1a, b). Importantly, hypoxia-induced HIF-1α stabilization was not altered by reoxygenation for up to 15 min post-incubation when cells remained on ice, excluding the possibility that HIF-1α would be degraded by oxygen exposure during the trophozoite detachment or protein extraction steps (Fig. S2).

After 1.5 and 4.5 h of Giardia infection, no change in HIF-1α mRNA expression was observed compared to uninfected control cells (Fig. 1a). In hypoxic conditions, HIF-1α mRNA expression was significantly upregulated after 1.5 h of Giardia infection, but no change was observed after 4.5 h (Fig. 1a). HIF-1α protein expression was significantly increased in normoxic conditions at both the 1.5- and 4.5-hour time points in Giardia-infected cells compared to control cells (Fig. 1b). No change in HIF-1α protein expression was observed at either time point in hypoxic infected cells compared to hypoxic uninfected controls (Fig. 1c). The longevity of the cellular hypoxic response was also assessed at 24 h post-infection. No change in HIF-1α protein expression was observed in Giardia-infected cells compared to uninfected control cells in either oxygenic condition (Fig. 1b, c). In conclusion, Giardia drives HIF-1α stabilization early on during infection, but this stabilization does not persist after 24 h of infection.

Fig. 1.

Fig. 1

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Giardia duodenalis drives HIF-1α stabilization in normoxic intestinal epithelial cells. Caco-2 intestinal epithelial cells were incubated for 1.5, 4.5, or 24 h in normoxia (Nx) (~ 21% O2) or hypoxia (Hx) (1% O2). a HIF-1a mRNA expression was measured using RT-qPCR and normalized to β-actin. Data are presented as fold change relative to uninfected control cells at each time point in box plots with min to max whiskers. HIF-1α protein expression from b normoxic and c hypoxic conditions was quantified by western blotting and normalized to a β-actin loading control. Data represents the relative optical density of bands in box plots with the median and min to max whiskers. Original blots can be seen in Fig. S7. C = control uninfected cells; G = cells infected with G. duodenalis isolate GS/M trophozoites (MOI 1:10). * Indicates p < 0.05, ** indicates p < 0.01. n = 4–6.

Giardia upregulates cellular stress HIF-1 target genes in normoxic intestinal epithelial cells

To determine whether HIF-1α stabilization drives a transcriptional response in Giardia-infected cells, the expression of known HIF-1 target stress-related genes (i.e., VEGFA, ANKRD37, and GADD45A) was assessed via RT-qPCR. VEGFA, a key HIF-1 target gene involved in angiogenesis, was significantly upregulated in infected cells in both normoxia (p < 0.05) and hypoxia (p < 0.01) after 4.5 h (Fig. 2a). Noticeably, VEGFA gene expression was greater in normoxic infected cells (5.9-fold) compared to hypoxic infected cells (1.7-fold). ANKRD37 gene expression was next assessed, a key HIF-1 target gene that regulates cellular autophagy during hypoxic stress. In normoxia, expression of ANKRD37 was increased after 1.5 h (p = 0.057) and 4.5 h in infected cells (p < 0.05) (Fig. 2b). No change in the expression of ANKRD37 was observed in infected hypoxic cells compared to uninfected controls at either time point (Fig. 2b). Lastly, expression of GADD45A, a HIF-1 target gene involved in cell cycle regulation, was significantly upregulated after 1.5 h (p < 0.01), but not 4.5 h in normoxic infected cells (Fig. 2c). Conversely, GADD45A expression was significantly upregulated in hypoxic infected cells after 4.5 h (p < 0.01), albeit to a lesser degree compared to the normoxic condition (Fig. 2c). These data suggest that HIF-1 target cell stress-related genes are activated in non-differentiated Caco-2 cells in normoxia upon Giardia infection, but the response is less profound and varies temporally in the hypoxic condition. Given HIF-1α is a key regulator of cellular glycolytic activity, we next investigated the expression of HIF-1 target glycolytic genes.

Fig. 2.

Fig. 2

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Giardia duodenalis upregulates HIF-1 target cellular stress genes in intestinal epithelial cells. Caco-2 intestinal epithelial cells were incubated for 1.5–4.5 h in normoxia (~ 21% O2) or hypoxia (1% O2). mRNA expression of (a) VEGFA, (b) ANKRD37, and (c) GADD45A was measured using RT-qPCR and normalized to β-actin. Data are presented as fold change relative to uninfected control cells at each time point in box plots with min to max whiskers. C = control uninfected cells; G = cells infected with G. duodenalis isolate GS/M trophozoites (MOI 1:10). * Indicates p < 0.05, ** indicates p < 0.01. n = 4–6.

Glycolytic HIF-1 target genes are activated upon Giardia infection

In hypoxia, cells become more glycolytic to maintain bioenergetic homeostasis. Gene expression of the first glycolytic enzyme, Hexokinase 2 (HK2), was significantly upregulated after 1.5 h (p < 0.01) and 4.5 h (p < 0.05) in normoxic Giardia-infected cells (Fig. 3a). Despite these transcriptional alterations, western blotting illustrated no changes in HK2 protein expression at either time point in normoxia (Fig. 3d). No alterations to gene or protein expression of HK2 were observed in the hypoxic infected cells compared to uninfected hypoxic cells (Fig. 3a, e). Gene expression of phosphoglycerate kinase 1 (PGK1), an intermediary glycolytic enzyme, was elevated in hypoxic infected cells only after 1.5 h (p < 0.05), while no change was detected in normoxic infected cells (Fig. 3b). Expression of LDHA, a gene encoding lactate dehydrogenase that catalyzes the final reaction of anaerobic glycolysis, was significantly elevated in normoxic Giardia-infected cells after 4.5 h (p < 0.05) (Fig. 3c). No change in LDHA expression was observed in hypoxic infected cells at either time point (Fig. 3e). After 24 h of infection, no changes to glycolytic gene expression were observed in normoxic infected cells (Fig. 3a-c). Alternatively, expression of HK2 (p < 0.01), PGK1 (p < 0.05), and LDHA (p < 0.01) was significantly downregulated in infected hypoxic cells after 24 h (Fig. 3a-c). In all, glycolytic HIF-1 target genes were upregulated when HIF-1α was stabilized in normoxic infected cells.

Fig. 3.

Fig. 3

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Giardia duodenalis upregulates HIF-1 target glycolytic genes in intestinal epithelial cells. Caco-2 intestinal epithelial cells were incubated for 1.5, 4.5, or 24 h in normoxia (Nx) (~ 21% O2) or hypoxia (Hx) (1% O2). mRNA expression of (a) HK2, (b) PGK1, and (c) LDHA was measured using RT-qPCR and normalized to β-actin. Data are presented as fold change relative to uninfected control cells at each time point in box plots with min to max whiskers. HK2 protein expression from the (d) normoxic or (e) hypoxic conditions was quantified by western blotting and normalized to a β-actin loading control. Data represents the relative optical density of bands in box plots with min to max whiskers. Original blots can be seen in Fig. S8. C = control uninfected cells; G = cells infected with G. duodenalis isolate GS/M trophozoites (MOI 1:10). * Indicates p < 0.05, ** indicates p < 0.01. n = 4–6.

Giardia alters steady-state concentrations of intracellular glycolytic metabolites in intestinal epithelial cells

To further characterize the metabolic hypoxic signature upon Giardia infection, a metabolomic approach was used to investigate central carbon metabolism after 4.5 and 24 h of infection in normoxia and hypoxia. While no change was observed in intracellular glucose levels in both normoxic and hypoxic Giardia-infected cells, glucose-6-phosphate (G6P) was elevated in normoxic infected cells at both time points (p < 0.05) (Fig. 4a, b). We observed decreased pyruvate levels (p < 0.05) (Fig. 4h) and increased lactate levels (p < 0.05) (Fig. 4i) in normoxic infected cells after 4.5 h. Pyruvate and lactate are the precursor and product of lactate dehydrogenase, respectively. No other significant alterations to intracellular glycolytic intermediate metabolites were observed in infected cells in both normoxic or hypoxic cells after 4.5 h (Fig. 4c to h). After 24 h under normoxic conditions, we observed decreased levels of dihydroxyacetone phosphate (DHAP) (p < 0.01) (Fig. 4d) and phosphoenolpyruvic acid (PEP) (p < 0.05) (Fig. 4g). In hypoxic infected cells, multiple alterations to glycolytic intermediates were observed after 24 h, including decreased fructose-1,6-bisphosphate (p < 0.01), decreased DHAP (p < 0.05), and increased PEP (p < 0.05) (Fig. 4c, d, g). No change in intracellular lactate was observed in hypoxic infected cells (Fig. 4i). Measurement of extracellular lactate showed no significant differences between control and Giardia-infected cell spent media at all time points in normoxia or hypoxia (Fig. S3). While no significant alterations to Tricarboxylic Acid Cycle (TCA) metabolites were observed in normoxic infected cells at either time point, intracellular levels of citrate, α-ketoglutarate, and succinate were altered in hypoxic Giardia-infected cells (p < 0.05) (Fig. 5a, c, e). No significant alterations in fumarate or malate were observed in either condition at either time point (Fig. 5b, d). Significant alterations to other intracellular metabolites, including amino acids and nucleosides, were also observed in Giardia-infected cells in both normoxic and hypoxic conditions (Fig. S4, S5, S6). Overall, Giardia alters the steady-state cellular concentration of numerous glycolytic metabolites more prominently in normoxic cells after 4.5 h. However, hypoxia modulates the steady-state concentrations of both glycolytic and TCA metabolites in infected cells after 4.5 and 24 h of infection.

Fig. 4.

Fig. 4

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Giardia duodenalis alters intracellular glycolytic metabolites in intestinal epithelial cells. Caco-2 intestinal epithelial cells were incubated for 4.5–24 h in normoxia (~ 21% O2) or hypoxia (1% O2). Intracellular (a) glucose, (b) glucose-6-phosphate, (c) fructose 1,6-bisphosphate, (d) dihydroxyacetone phosphate, (e) glycerol-3-phosphate, (f) 3-phosphoglyceric acid, (g) phosphoenolpyruvic acid, (h) pyruvate, and (i) lactate were analyzed via LC-MS. Box plots with min to max whiskers represent the fold change (f.c.) averages relative to uninfected control averages of each metabolite per time point. C = control uninfected cells; G = cells infected with G. duodenalis isolate GS/M trophozoites (MOI 1:10). * Indicates p < 0.05, ** indicates p < 0.01 n = 5.

Fig. 5.

Fig. 5

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Giardia duodenalis alters intracellular TCA metabolites in hypoxic intestinal epithelial cells. Caco-2 intestinal epithelial cells were incubated for 4.5–24 h in normoxia (~ 21% O2) or hypoxia (1% O2). Intracellular (a) citrate, (b) fumarate, (c) α-ketoglutarate, (d) malate, and (e) succinate were analyzed via LC-MS. Box plots with min to max whiskers represent the fold change (f.c.) averages relative to uninfected control averages of each metabolite per time point. C = control uninfected cells; G = cells infected with G. duodenalis isolate GS/M trophozoites (MOI 1:10). * Indicates p < 0.05, ** indicates p < 0.01 n = 5.

Discussion

The gastrointestinal epithelium exists in a state of physiologic hypoxia that can be further exacerbated in the presence of pathogens or inflammation35–37. This study investigated the role of HIF-1α during G. duodenalis infection. The findings indicate that Giardia stabilizes HIF-1α protein in human intestinal epithelial cells in normoxic and hypoxic conditions. Giardia-dependent HIF-1α stabilization upregulates the transcription of HIF-1 target stress-related genes VEGFA, ANKRD37, and GADD45A. Moreover, the increased transcription of HIF-1 target glycolytic genes (HK2 and LDHA) and the altered steady-state concentrations of intracellular glycolytic metabolites (G6P, DHAP, PEP, pyruvate, and lactate) indicate that intestinal epithelial cells undergo a change in central carbon metabolism upon Giardia infection.

The regulation of small intestinal oxygen homeostasis in the context of Giardia infection is poorly understood. Duodenal epithelial cells infected with G. duodenalis strain WB demonstrated lower protein expression of ZO-2 and increased expression of occludin in anaerobic conditions38. This barrier dysfunction was determined to be a result of the dephosphorylation of protein kinase C by the parasite in the absence of oxygen, indicating oxygen availability can directly alter host-parasite interactions on the cellular level38. Transcriptomic studies have also suggested that Giardia is associated with a hypoxic signature in mammalian cells. Human organoid monolayers infected with G. duodenalis isolate WB illustrated enrichment of hypoxia-response-related genes (Molecular Signatures Database) after 24 h of infection31. Similarly, HIF-1 target genes (NOS2, ANKRD37, GADD45A, ITF, MIR210HG, and SLC2A3) were upregulated in Caco-2 monolayers infected with G. duodenalis isolates WB and GS/M after 4.5 h of infection32,34. Recently, it was found that Caco-2 cells exposed to Giardia extracellular vesicles (EVs) displayed transcriptional enrichment of the HIF-1 signaling pathway, suggesting that EVs alone may in part drive an epithelial hypoxic response33. Furthermore, Giardia excretory-secretory products from both Assemblage A and B isolates increased nuclear translocation of NFκB in Caco-2 cells despite decreases in ERK1/2 and P38 phosphorylation, suggesting hypoxic responses may play a role39. These findings may suggest that although the small intestine faces low concentrations of oxygen, the alterations induced by Giardia may alter local oxygen equilibrium and activate an epithelial hypoxic response.

While several reports suggest a hypoxic cellular response occurs during giardiasis, the stabilization of HIF-1α protein, the key cellular regulator of the hypoxic adaptation, has yet to be elucidated. This study aimed to characterize the role of HIF-1 and associated metabolic alterations in intestinal epithelial cells infected with Giardia. The results indicate that Giardia drives stabilization of HIF-1α under normoxic conditions, as well as an increase in the transcription of known HIF-1 target genes aimed at combating cellular stress. However, no change was observed in HIF-1α transcription despite the strong increase in protein expression in normoxic infected cells. This observation confirms the assertion that HIF-1α is constitutively transcribed at a high rate, while post-translational modifications are the true regulators of the subunit’s activity40. The profound increase in VEGFA expression observed in the normoxic Giardia-infected cells is consistent with the induction of a hypoxic response observed in numerous other protozoa, including Entamoeba histolytica, Leishmania major, Trypanosoma brucei, and Plasmodium berghei30,41–44. Giardia failed to increase the stabilization of HIF-1α in hypoxia, begging the question of whether this reflects a robust stabilization of HIF-1α that could not be detected at higher levels using Western blotting.

Increased expression of HIF-1 target genes was observed in normoxic Giardia-infected cells, however, hypoxic responses varied between 1.5- and 4.5-hours post-infection. Given that GADD45A is involved in the regulation of the cell cycle during cellular stress, it is possible that increased expression of this gene may function to reallocate cellular energy towards survival versus replication early in infection45. Alternatively, ANKRD37 expression is linked to the regulation of cellular autophagy which is often associated with later stages of infection and inflammation46. A previous study indicated that Giardia infection is associated with autophagy in Caco-2 cells after 6 h of infection, consistent with our observation of increased ANKRD37 expression after 4.5 h of infection47. Moreover, Giardia-driven autophagy was associated with mTOR signaling which can also drive HIF-1α stabilization, further strengthening this link between Giardia infection and hypoxic cellular responses47,48. Gene expression was not increased in infected-hypoxic cells to the degree it was in normoxic cells, consistent with the hypothesis that under normoxic conditions, the HIF-1α stabilization driven by Giardia infection was so significant that hypoxic conditions were only able to increase gene expression marginally during infection. More research is needed to better understand the temporal regulation of HIF-1 target genes and associated transcriptomic signatures upon Giardia infection in the context of hypoxia.

The present study found that Giardia infection was associated with increased transcription of genes encoding key glycolytic enzymes, consistent with previous findings that HIF-1 increases expression of glycolytic genes during infection29,49,50. This finding is also corroborated by transcriptomic data from differentiated Caco-2 cells infected with G. duodenalis isolate GS/M which revealed significant upregulation of HK2 and Phosphofructokinase (PFK) after 4.5 h of infection, and enrichment of genes related to the regulation of glycolytic processes34. Interestingly, when HK2 expression was elevated in normoxic infected cells, intracellular G6P was also elevated in this study. HK2 converts glucose to G6P and thereby traps carbon in the cell. Importantly, previous findings have indicated that increased glucose import by the sodium-dependent glucose cotransporter (SGLT1) via active transport is protective against Giardia-induced apoptosis of intestinal epithelial cells51. However, HIF-1α and hypoxic responses favour the activity of passive glucose transporters (i.e., GLUT family) which are reliably expressed in Caco-2 cells, while the SGLT1 transporter is not29,52–54. Together, these observations suggest that the increase in intracellular G6P observed in this study may be related to the previously reported cytoprotective role of glucose uptake during Giardia infection.

Similar to the trend observed with HK2 and G6P, the increase in LDHA expression observed at 4.5 h in normoxic infected cells was concomitant with an increase in its metabolic product, lactate. LDHA enables the anaerobic conversion of pyruvate to lactate, converting NADH into the oxidized NAD + cofactor needed for glycolysis. The Caco-2 intestinal epithelial cell line used in this study is known to secrete lactate, and other studies have shown that HIF-1α drives expression of LDHA and increased production of lactate in both cancerous and non-cancerous epithelial cells49,55. This study confirms these findings and shows that Giardia-infected intestinal epithelial cells have an induced hypoxic phenotype similar to the HIF-1 driven phenotype observed in the context of other protozoan infections43,56,57.

Hypoxia has a direct impact on the TCA cycle and these impacts can be further modulated by HIF-1α in various cancer cells58–60. As expected, TCA metabolites were significantly altered in hypoxic cells after 24 h of infection. However, the present findings revealed no alterations in TCA cycle metabolite levels despite the stabilization of HIF-1α in normoxic infected cells. Furthermore, after 24 h, the stabilization of HIF-1α in normoxic infected cells was lost and levels of glycolytic gene expression returned to normal. This observation may suggest that the Giardia-induced hypoxic-like response is transient. Interestingly, other Giardia-induced cellular responses have been observed early on during infection and do not persist, including altered ion secretion, increased transcription of inflammatory cytokines and immediate-early response genes31,61. Giardia also undergoes differential transcriptional alterations during infection and throughout it’s life cycle, but more research is needed to understand how changes to the Giardia transcriptome affect host responses62.

This study has highlighted a novel phenomenon wherein G. duodenalis isolate GS/M activates a hypoxic response in vitro in intestinal epithelial cells. The findings pave the way toward several future research directions. First, Caco-2 cells used in the present study are cancer cells, and hence, have a high glycolytic capacity. More research is warranted using primary epithelial cells. Furthermore, to better understand the physiological relevance of this observation, future studies should be carried out using both wild-type and HIF-1α knockout mice to uncover the role of HIF in vivo during giardiasis. Finally, post-translational modifications other than hydroxylation (i.e., phosphorylation, acetylation, sumoylation, lactylation) have been implicated in regulating HIF stability and localization. Investigation into these post-translational modifications of HIF-1α in giardiasis offers fertile ground for future research to better understand the regulation and intracellular trafficking of HIF-1a during intestinal infection.

In summary, we report a novel phenotype wherein Giardia induces a transient hypoxic-like response under normoxic conditions in intestinal epithelial cells. This phenotype is characterized by increased protein expression and transcription of HIF-1 target genes related to cell stress and central carbon metabolism. Recent research has started to investigate the cellular hypoxic response to G. duodenalis genetic Assemblage A (WB isolate)31–33. The present study complements these findings by using a Giardia genetic Assemblage B (GS/M) and sheds new light on the role of oxygen in the context of enteric infections. Future research is warranted to understand whether this observed phenomenon is specific to Assemblage B isolates and whether HIF-1α contributes to differences in virulence between G. duodenalis Assemblages.

Methods

Cell culture

This investigation utilized non-differentiated Caco-2 human intestinal epithelial cells (ATCC-HTB-37), a well-established model for the study of giardiasis, to determine the effect of the parasite on cellular hypoxic responses6,32,63,64. Cells were cultured between passages 20 and 35 with high glucose DMEM media (Gibco) containing 4.5 g/L of glucose, supplemented with 10% heat-inactivated FBS and 50 U/mL Penicillin-Streptomycin. Cells were seeded in 6-well plates (3 × 105 cells/mL) and grown to confluency (37 °C, 5% CO2).

Giardia duodenalis culture

All coculture experiments in this study utilized Giardia duodenalis isolate GS/M (Assemblage B) purchased from ATCC (Manassas, VA). Giardia trophozoites were grown axenically in conical polystyrene centrifuge tubes with Kiester’s modified TYI-1-S33 media, supplemented with 50 U/mL Penicillin-Streptomycin (Thermo Fisher Scientific)65. Giardia cultures were maintained in an incubator at 37 °C (5% CO2) and once confluent, tubes were placed on ice to detach trophozoites for passaging or in preparation for cocultures. Detached trophozoites were washed with ice-cold PBS, then counted and resuspended in warm DMEM media (Gibco) to achieve a concentration of 5 × 106 trophozoites per mL. In the 6-well plates, cells were treated with 2 mL of Giardia-containing media to achieve a multiplicity of infection (MOI) of 10 (1 × 107 trophozoites per well)6,66.

Normoxic and hypoxic coculture incubations

Confluent Caco-2 cells were washed with warm PBS before the addition of treatment media for cocultures. Coculture incubations took place in the same incubator at 37 °C for 1.5, 4.5, or 24 h. For the normoxic condition, cocultures were placed in the incubator exposed to atmospheric oxygen levels (~ 21% O2) and 5% CO2. For the hypoxic condition, cocultures were placed into a hypoxia chamber (Stemcell Technologies) immediately after preparation. The hypoxia chamber was connected to a gas tank containing a hypoxia mix (1% O2, 5% CO2, and balanced with N2) (Linde Canada). At the beginning of each hypoxic experiment, the chamber was flushed for 4 min at a flow rate of 20 L per minute per the manufacturer’s protocol to establish a hypoxic environment (Stemcell Technologies). For the 4.5-hour and 24-hour hypoxic cocultures, the tank was re-flushed every 1.5 and 3 h, respectively (Stemcell Technologies). Experimental validation of HIF-1α stabilization in hypoxic conditions in Caco-2 cells was conducted in an anaerobic glove box (Coy Laboratories, USA) equilibrated to 1% O2.

Cellular RNA extraction and quantitative RT-PCR analysis

After the coculture incubation, 6-well plates were placed on ice and wells were washed with cold PBS to promote trophozoite detachment. Cells were lysed with 600 µL of RLT Buffer (Qiagen) supplemented with 2-mercaptoethanol (10 µL per mL), and total RNA was extracted using the RNeasy Mini Kit (Qiagen) according to the manufacturer’s protocols. After quantifying the RNA concentrations using the Nanodrop, complementary DNA was synthesized using the QuantiTect Reverse Transcription (RT) Kit (Qiagen). Reverse-transcriptase quantitative polymerase chain reaction (RT-qPCR) was carried out using the QuantiTect SYBR Green kit (Qiagen) on a RotorGeneQ cycler (Qiagen). Previously published primer sequences were utilized to assess mRNA expression of HIF-1α and HIF-1 target genes (Table S1)67–73. Relative gene expression was calculated using the 2−ΔΔCt method, normalized to β-actin as a housekeeping gene. Data is presented as fold change to uninfected control averages.

Western blotting

Coculture experiments were repeated for optimized protein extraction based on previously published protocols49. Once trophozoites were detached as described above, cells were lysed on ice for 10 min with 200 µL of RIPA lysis buffer (Sigma Aldrich) supplemented with protease inhibitors (1 tablet per 10 mL) (Sigma Aldrich). Cell lysates were then scraped and transferred to microcentrifuge tubes, vortexed, and kept on ice for 20 min. Lysates were centrifuged for 15 min at 17,950 g. The protein concentration of each supernatant was quantified using a BCA protein assay kit (Thermo Scientific). Samples were diluted with ddH2O to achieve 20 µg in 15 µL of sample and were prepared for separation by SDS-polyacrylamide gel electrophoresis with sample buffer (5x Laemmli blue buffer and 5% 2-mercaptoethanol). Protein samples were placed in a heating block at 100 °C for 5 min and loaded into 15-well 1.5 mm gels (8% resolving, 4% stacking). Gels were run at 150 V for 10 min. Once samples had run beyond the stacking gel, gels were run at 275 V for 30 min or until the dye reached the bottom of the gel.

Nitrocellulose membranes (Amersham) were soaked in 1X cold transfer buffer for 10 min before the wet transfer (320 mA for 70 min). Membranes were incubated overnight at 4 °C with antibodies for HIF-1α (1:2500, #36169, Cell Signaling), HK2 (1:5000, #2867, Cell Signaling), or β-actin (1:5000, #4970, Cell Signaling) in 5% milk. The following day, membranes were washed thrice for 15 min with 1X TBS-T, followed by a one-hour incubation with anti-rabbit secondary antibody (1:10,000, #7074, Cell signaling) in 5% milk at room temperature. Membranes were washed again as per above and developed using 200 µL of ECL (Amersham) for visualization using a ChemiDoc (BioRad). Densitometric analysis was carried out using Image J. β-actin was used as a loading control to determine the relative optical density of bands of interest.

Characterization of intracellular metabolites

Coculture experiments were repeated to extract intracellular metabolites as previously described with optimizations74. After trophozoites were detached as described above, plates remained on ice and 1 mL of 90% methanol (0.5% formic acid) was added to each well for 20 min. After scraping the wells, samples were centrifuged at 10,620 g for 10 min (4 °C). Supernatants were placed in a SpeedVac for 12 h, and concentrated metabolites were then resuspended in 100 µL of 50% methanol. Samples were analyzed using liquid-chromatography mass-spectrometry (LC-MS) at the Calgary Metabolomics Research Facility (CMRF). Reverse-phase ion-pairing (RPIP) methods were used to quantify a panel of central carbon metabolites, as previously described (Table S2)75,76. Peak picking and data analysis were performed using El-Maven software.

Measurement of extracellular lactate

Extracellular release of lactate was measured in the spent media of Giardia-infected Caco-2 cells. 500 µL of media was collected from each well upon completion of the coculture incubation and centrifuged at maximum speed (20,817 g) for 10 min to pellet and remove planktonic trophozoites. Extracellular lactate was isolated and quantified following the lactate assay kit protocol (Cayman). Fluorescence readings of the assay plates were made using an excitation and emission wavelength of 540 nm and 595 nm, respectively.

Statistical analysis

GraphPad (GraphPad Software Inc., San Diego, CA, U.S.A.) was utilized for statistical analysis. The normality of data was assessed before statistical analysis. A comparison between two sets of parametric data with normal distribution was performed using a Student’s T-test. Data sets with nonparametric data were analyzed using a Mann-Whitney test. All data is presented as box plots (interquartile range in box) with the median and min to max whiskers. p < 0.05 (*) was deemed statistically significant, with the degree of significance indicated by the number of stars (**=p < 0.01, ***=p < 0.001, ****=p < 0.0001). For the metabolomics dataset, a Benjamini-Hochberg adjustment was applied to account for multiple comparisons using a false discovery rate of 1%.

Supplementary Information

Below is the link to the electronic supplementary material.

Supplementary Material 1 (3.3MB, docx)

Acknowledgements

Funding for this project was provided by the Natural Sciences and Engineering Research Council of Canada (NSERC RT690446). E.D. was financed by Alberta Graduate Excellence Scholarships, as well as the Natural Sciences and Engineering Research Council of Canada- Canadian Graduate Students Master’s Scholarship. Metabolomics data were acquired at the Calgary Metabolomics Research Facility (CMRF), which is part of the Alberta Centre for Advanced Diagnostics (ACAD; PrairiesCan RIE #22734) and is supported by the International Microbiome Centre and the Canada Foundation for Innovation (CFI-JELF 34986).

Author contributions

T.A., A.B and E.D. conceived and designed the study. E.D., O.S. performed the in vitro assays, RT-qPCR, western blots, and lactate assay. I.L. provided metabolomic resources and discussed metabolomic results. C.T.T. and D.F. helped designing Western Blot experiments. E.D., T.A. and A.B discussed the results and wrote the manuscript with contributions from all authors.

Data availability

The datasets generated during this study are available from the corresponding author on reasonable request.

Declarations

Competing interests

The authors declare no competing interests.

Footnotes

Publisher’s note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.

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Associated Data

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Supplementary Materials

Supplementary Material 1 (3.3MB, docx)

Data Availability Statement

The datasets generated during this study are available from the corresponding author on reasonable request.

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