In aerobic organisms, the dioxygen molecule (O₂) is essential for many biochemical pathways, particularly as the final electron acceptor for bioenergetics. Hypoxia—conditions of decreased O₂ availability—is both an essential stimulus for normal development and a pathological trigger of cellular dysfunction and eventual cell death for humans and other mammals (Bickler and Buck, 2007). To maintain O₂ homeostasis, aerobic organisms have developed diverse cellular mechanisms for sensing and responding to alterations in O₂ level. For multiorgan organisms, the term ‘hypoxia’ is often loosely used to describe decreased O₂ levels. More precisely, the term ‘tissue hypoxia’ is meaningful when used in comparison to the baseline for the tissue. Physiological tissue O₂ (physoxia, the typical range of function), physiological hypoxia (reduction or fluctuation of pO₂ into a range at which adaptation is possible), and hypoxia with pathological impact (pO₂ at which cellular injury and death occur) are often cited as ~5, 2, and 1%, respectively, for humans (McKeown, 2014). However, these values can vary widely across tissues and even within a tissue and are affected by tissue-level regulation (e.g., blood flow) and cellular effects (e.g., changes in metabolic state) (Table 1; McKeown, 2014; Ortiz-Prado et al., 2019; Carreau et al., 2011; Jagannathan et al., 2016; Cigognini et al., 2016; Donovan et al., 2010; Mas-Bargues et al., 2019). Here, we focus on O₂ sensing in humans, using the term ‘hypoxia’ to denote decreased O₂ level relative to physoxia, that is, encompassing both physiological hypoxia and hypoxia with pathological impact.

Discovery of the PHD-HIF-pVHL pathway was pivotal to understanding hypoxia responses and has been reviewed extensively (Majmundar et al., 2010; Kaelin and Ratcliffe, 2008; Ivan and Kaelin, 2017; Schofield and Ratcliffe, 2004). Briefly, in normoxia, prolyl hydroxylase domain proteins (PHDs) use O₂ as a substrate to hydroxylate prolines on the transcription factor hypoxia-inducible factor α subunit (HIFα, i.e., HIF1α or HIF2α). The hydroxylated form of HIFα is recognized by the E3 ubiquitin ligase pVHL (von Hippel-Lindau protein), which promotes degradation of HIFα. By contrast, in hypoxia, the decreased catalytic activity of PHDs results in decreased hydroxylation and hence decreased pVHL recognition of HIFα, promoting the accumulation of HIFα. HIFα then translocates to the nucleus and, as a heterodimer with HIF1β, regulates transcription of a broad range of target genes. Thus, PHDs directly sense a decrease in the availability of molecular O₂ and transduce this signal to downstream effectors.

What defines a molecular hypoxia sensor? In engineering, a sensor is a device that detects changes to a physical property and transmits this information so that a system can respond to this change. Here, by analogy to human-engineered sensors, we define biological hypoxia sensors as proteins that (1) directly interact with O₂ molecules, (2) have activities that are strongly affected by physiological hypoxia, and (3) are coupled to downstream responses that depend on changes of their activities. Of note, many proteins respond to hypoxia by acting downstream of a sensor (e.g., HIF acting downstream of PHD) or by responding to changes in the cellular redox state. In this review, we specifically exclude these as they are not direct sensors of molecular O₂ —that is, their response to changes in O₂ levels does not involve direct interaction with O₂ (Bickler and Donohoe, 2002).

Strong candidates for hypoxia sensors include O₂-dependent enzymes, which by definition meet criterion 1. These enzymes constitute a mechanistically, structurally, and biologically diverse group of proteins. There are a number of reviews on the enzymology (Islam et al., 2018; Palfey and McDonald, 2010; Decker and Solomon, 2005; Jasniewski and Que, 2018; Biringer, 2020; Ferguson-Miller and Babcock, 1996; Finney et al., 2014; Bassan et al., 2003; Guengerich, 2007; Ponnaluri et al., 2013; Ivanov et al., 2010; Daff, 2010; Wikström et al., 2018; Huang and Groves, 2018; Romero et al., 2018; Sono et al., 1996; Martinez and Hausinger, 2015; Roberts and Fitzpatrick, 2013; Itoh, 2006; Solomon et al., 2001; Bugg, 2001; Abu-Omar et al., 2005), biological function (Schofield and Ratcliffe, 2004; Islam et al., 2018; Paton and Ntambi, 2009; Shmakova et al., 2014; Danielson, 2002; Donkó et al., 2005; Bundred et al., 2018; Fletcher and Coleman, 2020; Kuhn et al., 2015; Zhuang et al., 2015; Kooistra and Helin, 2012; Mashima and Okuyama, 2015; Wu and Zhang, 2017; Johansson et al., 2014; Daubner et al., 2011; Fong and Takeda, 2008; Markolovic et al., 2015), and evolution (Danielson, 2002; Taylor and McElwain, 2010; Chandrasekharan and Simmons, 2004; Wilks, 2002) of individual subclasses of O₂-dependent enzymes. Here, we provide a global map of human O₂-dependent enzymes in potential hypoxia sensing. We first survey the broad categories and then discuss specific members that are known or speculated hypoxia sensors. Finally, we investigate the links between O₂-dependent enzymes and hypoxia-related evolutionary adaptations and diseases.

O₂-dependent enzymes as hypoxia sensor candidates

We start by providing background and taxonomies for considering the three basic ‘sensor’ requirements discussed above.

First, O₂-dependent enzymes directly interact with O₂ molecules as one of the substrates. Non-enzymatic proteins that directly interact with O₂, for example, globins, have been reviewed elsewhere (Burmester and Hankeln, 2014; Lecomte et al., 2005). In humans, 221 enzymes are known or likely to be O₂-dependent (Supplementary file 1), that is, utilizing O₂ as an electron acceptor for the oxidation of other substrates. Based on their catalyzed reactions, O₂-dependent enzymes can be divided into three subclasses: dioxygenases, which catalyze the insertion of both oxygen atoms of the O₂ molecule into substrates; monooxygenases, which catalyze the insertion of one oxygen atom of the O₂ molecule into a substrate and the reduction of the other oxygen atom to H₂O; and oxidases, which catalyze the reduction of O₂ molecules to 2H₂O or H₂O₂ (Figure 1).

Figure 1

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Second, O₂-dependent enzymes have diverse mechanisms for utilizing O₂ as a substrate, resulting in different sensitivities to O₂ concentrations. Sensitivity is determined, in part, by the binding affinity of O₂ with the enzyme’s catalytic center. Most O₂-dependent enzymes (177/221) use, or are speculated to use, O₂-binding metal ions at their catalytic centers. Factors affecting the O₂-binding affinity include the metal center (iron or copper in humans), ligands (enzyme residues and other substrates) for the metal center, and the environment of the catalytic pocket. The other O₂-dependent enzymes with known non-metal catalytic centers (37/221) utilize flavin adenine dinucleotide (FAD) or flavin mononucleotide (FMN) to activate O₂. For these enzymes, the accessibility of O₂ to FAD or FMN at the catalytic center affects the binding affinity. Dioxygenases, monooxygenases, and oxidases can be further subdivided by their catalytic centers (Table 2). Ultimately, these mechanisms affect the threshold at which enzymatic activities are saturated with O₂, thus determining whether the enzyme’s activities are strongly affected by physiological-range hypoxia.

Table 2

Category	Subcategory by catalytic center	Metal species at catalytic center	Ligands for the metal species at catalytic center (cofactor/substrate and enzyme residues)	Number of enzymes
Dioxygenase	2-OG-dependent dioxygenase	Fe	2-OG, His, His, Asp/Glu	59
	Heme-dependent dioxygenase	Fe	Heme, His	5
	Lipoxygenase	Fe	His, His, His, Ile, His/Asa/Asn/none	6
	Others	Fe	His, His, His/Asp/Glu*	10
Monooxygenase	Heme-dependent monooxygenase	Fe	Heme, Cys/His/Glu	61
	Non-Heme Fe-dependent monooxygenase	Fe	His, His, His/Asp/Glu*	9
	Cu-dependent monooxygenase	Cu	His, His, Met	5
	Flavin-dependent monooxygenase	None (uses flavin)	N/A	12
	Others†	N/A	N/A	2
Oxidase	Heme-copper	Fe and Cu	His, His, His for Cu; Heme and His for Fe	1
	Fe-dependent oxidase	Fe	Varies	14
	Cu-dependent oxidase	Cu	Varies	7
	Flavin-dependent oxidase	None (uses flavin)	N/A	25
	Others†	N/A	N/A	5

1. *

   Substrates/cofactor ligands for this category varies for each member depending on the reaction it catalyzes.

2. †

   Members in this category are not fully studied.

Third, O₂-dependent enzymes regulate diverse cellular processes: (1) oxidative phosphorylation is responsible for mitochondrial ATP production and cellular survival (Hatefi, 1985); (2) post-translational modifications (hydroxylation, demethylation, or thiol oxidation) of proteins can regulate protein conformation, stability, and activity (Schofield and Ratcliffe, 2004; Bundred et al., 2018; Fletcher and Coleman, 2020; Kooistra and Helin, 2012; Johansson et al., 2014); (3) hydroxylation and demethylation of DNA/RNA can regulate DNA damage repair, epigenetic modifications, and transcription/translation (Wu and Zhang, 2017; Zaccara et al., 2019); (4) metabolism of amino acids and lipids can maintain cellular hemostasis and regulate cellular pathways through signaling molecules (Paton and Ntambi, 2009; Kuhn et al., 2015; Mashima and Okuyama, 2015; Daubner et al., 2011; Chandrasekharan and Simmons, 2004; Ball et al., 2014); and (5) metabolism of xenobiotics can regulate drug clearance and detoxification (Danielson, 2002; Poulos and Johnson, 2005). Typically, dioxygenases have macromolecules as substrates and regulate cellular processes at a transcriptional or translational level (Schofield and Ratcliffe, 2004; Islam et al., 2018; Bundred et al., 2018; Fletcher and Coleman, 2020; Kooistra and Helin, 2012; Wu and Zhang, 2017; Johansson et al., 2014; Hancock et al., 2015), while monooxygenases and oxidases often have small molecules as substrates and function in metabolism (Romero et al., 2018; Paton and Ntambi, 2009; Danielson, 2002; Daubner et al., 2011). Together, O₂-dependent enzymes are integral to a plethora of physiological processes in aerobic animals.

Candidate hypoxia sensors can be identified among the O₂-dependent enzymes, in part by the binding affinity between O₂ and the enzyme as quantified by the O₂ K_m value, which suggests the level at which the enzyme is most sensitive to changes in O₂ (Kaelin and Ratcliffe, 2008; Schofield and Ratcliffe, 2004; Shmakova et al., 2014; Hancock et al., 2015; Wilson et al., 2020; Holdsworth and Gibbs, 2020; Baik and Jain, 2020). (The Km value, also known as the Michaelis constant, is the concentration of a substrate at which an enzymatic reaction rate is 50% of the maximum reaction rate. A larger K_m value reflects lower O₂ affinity.) Importantly, the measured K_m value is affected by the measurement method, for example, mass spectrometry vs. isotope assays. Besides the Km value, other cellular factors such as the concentration and conformation of the enzyme, as well as concentrations of other substrates or products, also affect the net enzymatic activity and hence the downstream effects of the enzyme. Beyond cellular-level effects, whether an O₂-dependent enzyme functions as a hypoxia sensor in vivo can depend on the tissue pO₂ context (Table 1). Taken together, whether an O₂-dependent enzyme functions as a hypoxia sensor in vivo depends not only on the O₂ K_m value but also on multiple other factors.

O₂-dependent enzymes that are known or potential hypoxia sensors

Below, we classify dioxygenases, monooxygenases, and oxidases into different subgroups based on their catalytic centers and discuss known (Figure 2A) and potential (Figure 2B) hypoxia sensors in each subgroup.

Figure 2

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Dioxygenases

Based on their catalytic centers, the dioxygenase family members can be further classified into 2-OG-dependent dioxygenase, heme-dependent dioxygenases, lipoxygenases, and other dioxygenases (Table 2).

2-Oxyglutarate (2-OG)-dependent dioxygenases

In humans, there are ~60 identified or postulated dioxygenases (Table 2, Supplementary file 1) that use 2-OG as the co-substrate to catalyze the hydroxylation of their primary substrates, which include proteins, nucleic acids, and lipids (Figure 3A, Figure 3—figure supplement 1; Islam et al., 2018; Fletcher and Coleman, 2020; Rose et al., 2011). We note that when hydroxylation occurs on the carbon of an N-methyl group, this can lead to demethylation, which occurs through spontaneous fragmentation to formaldehyde and the demethylated product (Figure 3B; Islam et al., 2018; Fletcher and Coleman, 2020; Rose et al., 2011).

Figure 3 with 3 supplements see all

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Among O₂-dependent enzymes, 2-OG-dependent dioxygenases are relatively well studied. A majority of members in this subgroup catalyze hydroxylation or demethylation on proteins, DNA, and RNA, and are involved in the regulation of transcription and translation (Islam et al., 2018; Fletcher and Coleman, 2020; Rose et al., 2011). We focus on three subgroups relevant to hypoxia biology: direct HIF modulators, epigenetic modulators, and translational modulators. These subgroups encompass most known hypoxia sensors, including PHDs, factor inhibiting HIF (FIH1), lysine demethylases (KDMs), and ten-eleven translocation methylcytosine dioxygenases (TET1-3), as well as potential sensors that have impaired activities during hypoxia. For each subgroup, we highlight the most well-known sensors and propose additional, potential sensors.

Direct HIF modulators

These include PHDs (which catalyze prolyl hydroxylation of HIFα) and FIH (which catalyzes asparaginyl hydroxylation of HIFα) (Table 3).

Table 3

Gene symbol	Protein name	Type of reaction	Hydroxylation sites in HIFα	Non-HIF substrate examples
EGLN1	PHD2	Prolyl hydroxylation	HIF1α Pro402, Pro564; HIF2α Pro405, Pro531; HIF3α Pro492	FLNA, Akt
EGLN2	PHD1	Prolyl hydroxylation	HIF1α Pro402, Pro564; HIF2α Pro405, Pro531; HIF3α Pro492	FOXO3, Cep192, TP53
EGLN3	PHD3	Prolyl hydroxylation	HIF1α Pro564; HIF2α Pro405, Pro531; HIF3α Pro492	ATF-4, ADRB2, TP53
HIF1AN	FIH1	Asparaginyl hydroxylation	HIF1α Asn803, HIF2α Asn847	IκBα, Notch, OTUB1, RIPK4

The PHD enzymes and their critical role in regulating the PHD–HIF-pVHL signaling pathway are a paradigm for cellular sensing and response to hypoxia (Figure 2A; Majmundar et al., 2010; Kaelin and Ratcliffe, 2008; Ivan and Kaelin, 2017; Schofield and Ratcliffe, 2004). In humans, HIF is composed of an α subunit (HIF1α, HIF2α, or HIF3α) and invariant β subunit (HIF1β), and there are three PHD isoforms, namely PHD1 (EGLN2), PHD2 (EGLN1), and PHD3 (EGLN3). These PHDs are canonical sensors that illustrate our criteria for O₂ sensors.

First, PHDs directly interact with O₂, utilizing O₂ to hydroxylate prolines in the O₂-dependent degradation domain (ODD) of HIFα (Epstein et al., 2001; Hirsilä et al., 2003).

Second, the enzymatic activities of PHDs are sensitive to cellular/tissue hypoxia. The O₂-binding affinities of all three PHDs, represented by O₂ Km values, have been measured in vitro with HIF1α peptides as substrates. Using a short 19-residue HIF1α fragment as the substrate, the reported O₂ Km values for PHD1-3 are in the range of 229–1746 μM (Table 4; Hirsilä et al., 2003; Dao et al., 2009; Tarhonskaya et al., 2014). However, recent measurements using longer HIF1α fragments estimate O₂ Km values for PHD2 in the range of 67–85 μM (Table 4; Ehrismann et al., 2007), corresponding to pO₂ values in the (physoxia) range of 6–8% (Table 1), consistent with the sensitivities of PHDs to changes in physiological O₂ concentrations.

Table 4

Category	Enzyme*	Km for O2	Assay details	Reference
Dioxygenase	PHD2 (EGLN1)	250 μM	In vitro radioactivity 2-OG turnover assay with HIF1α (556–574) peptide as substrate	Hirsilä et al., 2003
		1746 μM	In vitro time-resolved fluorescence resonance energy transfer assay with P564-HIF1α peptide (DLEMLAPYIPMDDDFQL) as substrate	Dao et al., 2009
		67 μM	In vitro O2 consumption assay with HIF1α (502–697) peptide as substrate	Ehrismann et al., 2007
		81 μM	In vitro O2 consumption assay with HIF1α (530–698) peptide as substrate	Ehrismann et al., 2007
	PHD1 (EGLN2)	230 μM	In vitro radioactivity 2-OG turnover assay with HIF1α (556–574) peptide as substrate	Hirsilä et al., 2003
	PHD3	230 μM	In vitro radioactivity 2-OG turnover assay with HIF1α (556–574) peptide as substrate	Hirsilä et al., 2003
	KDM4E	197 μM	In vitro O2 consumption assay with ARK(me3)STGGK peptide as substrate	Cascella and Mirica, 2012
	KDM4A	173 μM	In vitro MALDI-TOF-MS assay with H31−15K9me3 peptide as substrate	Hancock et al., 2017
		57 μM	In vitro O2 consumption assay with ARK(me3)STGGK peptide substrate	Cascella and Mirica, 2012
		60 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–19)K9me3 as substrate	Chakraborty et al., 2019
	KDM6A	180 μM	In vitro radioactivity 2-OG turnover assay with histone H3(21–44)K27(me3) as substrate	Chakraborty et al., 2019
	KDM4C	158 μM	In vitro O2 consumption assay with ARK(me3)STGGK peptide substrate	Cascella and Mirica, 2012
	KDM4B	150 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–19)K9me3 as substrate	Chakraborty et al., 2019
	FIH	90 μM	In vitro radioactivity 2-OG turnover assay with HIF1α (788–822) peptide as substrate	Koivunen et al., 2004
	KDM5A	90 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–21)K4me3 as substrate	Chakraborty et al., 2019
	KDM3A	75 μM (7.6% O2) †	In vitro demethylation-formaldehyde dehydrogenase-coupled reaction assay with K224-monomethylated PGC-1α peptide as substrate	Qian et al., 2019
	KDM5B	40 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–21)K4me3 as substrate	Chakraborty et al., 2019
	P4HA1	40 μM	Standard P4H activity assay with (Pro-Pro-Gly)10 (Peptide Institute) as a substrate	Hirsilä et al., 2003
	KDM5C	35 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–21)K4me3 as substrate	Chakraborty et al., 2019
	TET1	30 μM	In vitro radioactivity 2-OG turnover assay with oligonucleotides containing a 5-mC as substrate	Laukka et al., 2016
		3.0 μM (0.31% O2) †	In vitro DNA hydroxymethylation assay with genomic DNA as substrate	Thienpont et al., 2016
	TET2	30 μM	In vitro radioactivity 2-OG turnover assay with oligonucleotides containing a 5-mC as substrate	Laukka et al., 2016
		5.2 μM (0.53% O2) *	In vitro DNA hydroxymethylation assay with genomic DNA as substrate	Thienpont et al., 2016
	KDM5D	25 μM	In vitro radioactivity 2-OG turnover assay with histone H3(1–21)K4me3 as substrate	Chakraborty et al., 2019
	KDM6B	20 μM	In vitro radioactivity 2-OG turnover assay with histone H3(21–44)K27(me3) as substrate	Chakraborty et al., 2019
	IDO1	11.5–24 μM	In vitro O2 consumption assay with L-Trp as substrate	Kolawole et al., 2015
	PTGS1	10 μM (sheep)	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	PTGS2	13 μM (mouse)	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	ALOX5	13 μM (porcine)	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	ALOX12	13 μM	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	ALOX15	26 μM (porcrine)	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	ALOX15	26 μM (rabbit)	In vitro radioactivity label assay with [1-14C]arachidonic acid as substrate	Juránek et al., 1999
	ADO	>500 μM	In vitro UPLC-MS-TOF assay with RGS4(2–15) peptide as substrate	Masson et al., 2019
Monooxygenase	NOS1 (nNOS)	350 μM (rat)	In vitro heme-NO complex formation assay with L-arginine as substrate	Abu-Soud et al., 1996
	NOS2 (iNOS)	130 μM (mouse)	In vitro heme-NO complex formation assay with L-arginine as substrate	Abu-Soud et al., 2001
	NOS3 (eNOS)	4 μM (bovine)	In vitro heme-NO complex formation assay with L-arginine as substrate	Abu-Soud et al., 2000
		25 μM (bovine)	In vitro heme-NO complex formation assay with N-hydroxy-L-arginine as substrate	Abu-Soud et al., 2000
	TH	16.2 μM (low-activity state); 46.1 μM (high- activity state);	In vitro radioactivity label assay with 3H-tyrosine as substrate	Rostrup et al., 2008
		12.6–26.7 μM (low-activity state); 28.8–42.9 μM (high-activity state)‡;	In vitro oxygraphic assay with tyrosine as substrate	Rostrup et al., 2008
		2.6–3.9 μM (2–3 mmHg, rat) *	In vitro radioactivity label assay with 3H-tyrosine as substrate	Katz, 1980
	TPH1	3.9~12.9 μM (3–10 mmHg, rat) †	In vitro radioactivity label assay with 3H-tryptophan as substrate	Katz, 1980
	PAH	17 μM	In vitro oxygraphic assay with phenylalanine as substrate	Rostrup et al., 2008
	PAM	70 μM (rat)	In vitro radioactivity label assay with [α-2H2]-N-acylglycine of different chain length as substrates	McIntyre et al., 2010
Oxidase	Cytochrome c oxidase	<0.1 μM (rat)	In vitro O2 consumption assay measuring O2 consumption of purified rat mitochondria at low phosphate potential ([ATP]/[ADP]*[Pi])	Bienfait et al., 1975
		1–3 μM (rat)	In vitro O2 consumption assay measuring O2 consumption of purified rat mitochondria at high phosphate potential	Bienfait et al., 1975
		0.5 μM (mouse)	Cellular assay measuring the ‘apparent K (m)’ for O2 or p 50 of respiration in 32D cells using high-resolution respirometry	Scandurra and Gnaiger, 2010
	AOC3	38 μM	In vitro enzymatic assay using purified human AOC3	Shen et al., 2012

1. *

   O₂-dependent enzymes that are known sensors are highlighted in bold; that are reported to be inhibited under hypoxia are highlighted in a light orange background; that are reported to be associated with positive selections in high-altitude populations are highlighted in red (also see Table 6).

2. †

   Km of these enzyme were reported with units as % O₂ or mmHg, and calculated according to Mas-Bargues et al., 2019; Place et al., 2017.

3. ‡

   Combined data for TH1/3/4 splicing isoforms.

Third, the decreased activity of PHDs during hypoxia triggers specific downstream responses (Figure 2A; Majmundar et al., 2010; Kaelin and Ratcliffe, 2008; Ivan and Kaelin, 2017; Schofield and Ratcliffe, 2004). Under normoxia, hydroxylated HIFα is recognized and polyubiquitinylated by the E3 ubiquitin ligase von Hippel-Lindau protein (pVHL), which then leads to proteasome-mediated degradation of HIFα. Under hypoxia, decreased O₂ concentration suppresses the activity of PHDs. This allows HIFα to accumulate and translocate to nucleus, where it associates with the constitutively expressed HIF1β and forms the heterodimer transcriptional factor HIF. HIF then recruits transcriptional co-activators p300 and CREP-binding protein (CBP), binds with hypoxia-responsive elements (HREs) on DNA, and subsequently activates its target genes. Products of HIF-regulated genes are involved in multiple cellular and systematic adaptations to hypoxia, including metabolic shift from OXPHOS to glycolysis, redox homeostasis, angiogenesis, and erythropoiesis.

Although all PHD isoforms have similar O₂ affinities, their different expression patterns and substrate preferences among the HIFα isoforms lead to differential regulation of hypoxia sensing and response by the PHD-HIF-pVHL pathway. Of the three PHD isoforms, PHD2 exerts the greatest control over the PHD-HIF-VHL pathway response to hypoxia (Berra et al., 2003) and it is ubiquitously expressed across mouse tissues and is the most abundant isoform (Appelhoff et al., 2004; Willam et al., 2006). PHD1 and PHD3 are more tissue-specific, with PHD1 most expressed in testis and PHD3 in heart (Willam et al., 2006). PHD2 favors HIF1α as substrate, while PHD1 and PHD3 favor HIF2α (Table 3; Appelhoff et al., 2004). Interestingly, PHD2 and PHD3 are themselves HIF target genes and can be induced during hypoxia to provide feedback regulation for the PHD-HIF-pVHL pathway (Epstein et al., 2001). The nonoverlapping and complex roles of different PHD isoforms are evidenced by knockout mice: Phd2-/- mice are embryonic lethal due to placental and heart defects, while postnatal whole-body knockout of Phd2 leads to polycythemia, increased angiogenesis, and heart defects (Takeda et al., 2006; Minamishima et al., 2008). Tissue-specific knockout of Phd2 in mouse heart or brain is protective against ischemic cardiac or neural injury, respectively (Kunze et al., 2012; Hölscher et al., 2011). Phd1 knockout mice have altered metabolism in skeletal muscles and overall enhanced hypoxia tolerance (Aragonés et al., 2008); and germline Phd3 knockout mice are hypotensive due to a hypofunctional sympathoadrenal system (Bishop et al., 2008).

There has been interest to identify non-HIF substrates for PHDs and other pathways that are regulated through PHD-catalyzed hydroxylation. In the last two decades, more than 20 non-HIF substrates for PHDs have been reported, and hypoxia-mediated hydroxylation of these proteins alters response of downstream pathways (Cockman et al., 2019; Strowitzki et al., 2019). Most of these substrates were identified through cellular studies, suggesting that PHDs may act upon non-HIF substrates under physiological conditions (Strowitzki et al., 2019; Zheng et al., 2014; Moser et al., 2013; Segura et al., 2016; Guo et al., 2016; Köditz et al., 2007; Xie et al., 2009; Ullah et al., 2017; Deschoemaeker et al., 2015; Rodriguez et al., 2018). One group found in in vitro enzymatic assays that PHDs lacked detectable activities on these non-HIF substrates. This difference in findings between cellular and in vitro enzymatic assays suggests that the action of PHDs on non-HIF substrates requires additional cellular machinery, such as adaptors or post-translational modifications (Cockman et al., 2019).

FIH1 is another 2-OG dioxygenase that is known to sense hypoxia and regulate the HIF pathway (Figure 2A; Lando et al., 2002a; Mahon et al., 2001). Under normoxia, FIH1 catalyzes the asparaginyl hydroxylation of the C-terminal transactivation domain (CTAD) of HIFα (Table 3; Koivunen et al., 2004; Lando et al., 2002b), which is responsible for its binding with the transcriptional coactivator p300/CBP (Lando et al., 2002b; Freedman et al., 2002). FIH1-catalyzed asparaginyl hydroxylation of the CTAD impairs the recruitment of p300/CBP and reduces transcriptional activity of HIF (Lando et al., 2002a; Lando et al., 2002b). In hypoxia, FIH1 activity is also reduced by hypoxia, enabling HIFα to recruit p300/CBP for transcriptional activation of its target genes (Lando et al., 2002a; Lando et al., 2002b). The reported O₂ Km value for FIHs is 90 μM, using a HIF1α peptide containing site Asn803 (Table 4; Koivunen et al., 2004). Compared with the Km values of PHDs in similar assays, FIH1 appears to be less sensitive to hypoxia, that is, as O₂ levels decrease, PHDs are inhibited before FIH1 (Tian et al., 2011). Thus, FIH1 is considered a fine modulator of the HIF pathway in sensing severe hypoxia. Consistent with the notion that its role is more limited, FIH1 knockout mice have abnormal metabolism but not other HIF-regulated processes (Zhang et al., 2010; Sim et al., 2018). There are also non-HIF substrates identified for FIH1 that may also be regulated in an O₂-dependent manner (Table 3; Cockman et al., 2009b; Scholz et al., 2016; Cockman et al., 2009a).

Epigenetic modulators

These include the lysine demethylase (KDM) Jumonji C (JmjC) domain-containing proteins, and the DNA demethylases ten-eleven translocation enzymes (TETs).

JmjC domain-containing proteins contain domains for Fe(II) and 2-OG binding and catalytic activities (Shmakova et al., 2014; Kooistra and Helin, 2012). They also bind with O₂ and utilize it as a substrate, therefore having the potential to function as hypoxia sensors if their O₂-binding affinities allow (Shmakova et al., 2014). Of the 32 identified JmjC proteins in humans, at least 23 conduct lysine demethylation reactions (Shmakova et al., 2014). Their substrates include both histone lysines (K4, K9, K27, and K36 on histone 3) and some non-histone lysines. (Table 5). Histone methylations affect chromatin structure and compactness and consequently regulate gene expression in either activating or silencing mode (Table 5; Shmakova et al., 2014; Kooistra and Helin, 2012; Walport et al., 2016; Li et al., 2022). H3K4me2/3, H3K9me2, H3K27me3, and H3K36me3 levels increase after hypoxia, possibly due to KDMs acting as O₂ sensors and effecting chromatin changes.

Table 5

KDM class	Members (gene symbol)	Histone lysyl residue substrates	Other substrates
KDM2	KDM2A	H3K36me1/me2 (A)	p65, NF-κB
	KDM2B	H3K36me1/me2 (A), H3K4me3 (A)
KDM3	KDM3A	H3K9me1/me2 (S)	PGC-1α K224me
	KDM3B	H3K9me1/me2 (S)
	JJMJD1C	H3K9me1/me2 (S)
KDM4	KDM4A	H3K9me2/me3 (S), H3K36me2 (A), H1.4K26me2/me3	WIZ, CDYL1, CSB, and G9a
	KDM4B	H3K9me2/me3 (S), H3K36me2 (A), H1.4K26me2/me3	WIZ, CDYL1, CSB, and G9a
	KDM4C	H3K9me2/me3 (S), H3K36me2 (A), H1.4K26me2/me3	WIZ, CDYL1, CSB, and G9a
	KDM4D	H3K9me2/me3 (S)
	KDM4E	H3K9me3 (S)	H3R2me2/me1, H3R8me2/me1, H3R26me2/me1, H4R3me2
KDM5	KDM5A	H3K4me2/me3 (A)
	KDM5B	H3K4me2/me3 (A)
	KDM5C	H3K4me2/me3 (A)	H3R2me2/me1, H3R8me2, H4R3me2a, ULK1R170me2a
	KDM5D	H3K4me2/me3 (A)
KDM6	KDM6A	H3K27me2/me3 (S)
	KDM6B	H3K27me2/me3 (S)
	KDM6C
KDM7	KDM7A	H3K9me1/me2 (S), H3K27me1/me2 (S)
	PHF8	H3K27me1/me2 (S), H4K20me1
	PHF2	H3K9me2/me3 (S)
Jmjc domain only	NO66	H3K4me2/me3 (A), H3K36me2/me3 (A)	Rpl8
	MINA53	H3K9me3 (S)	Rpl27a
	KDM8	H3K36me2 (A)	NFATc1
	JMJD6		H3R2me2,H4R3me2/me1, U2AF2/U2AF65, LUC7L2

1. *

   Known hydroxylation/demethylation sites are indicated.

KDM6A, also known as UTX, catalyzes demethylation at H3K27me2/me3 (Table 5; Hong et al., 2007). In 2019, Chakraborty et al. reported that increase in H3K27me3 levels during hypoxia is HIF-independent and is caused by the direct inhibition of KDM6A due to decreased pO₂ under hypoxia (Figure 2A; Chakraborty et al., 2019). The O₂ sensitivity of KDM6A was further confirmed by its O₂ Km value of 180 μM (Table 4), in a similar range as the PHDs and FIH and the highest among KDM6 members (Chakraborty et al., 2019). Sensing of hypoxia by KDM6A can control cell fate by chromatin reprogramming (Chakraborty et al., 2019). For example, it is reported that in mouse myoblast C2C12 cells, increase in H3K27me3 levels due to inactivation of KDM6A represses the expression of myogenic genes and blocks myogenic differentiation (Chakraborty et al., 2019).

KDM5A catalyzes demethylation at H3K4me2/me3 (Table 5) and was recently reported by Batie et al. to be another hypoxia sensor that could directly regulate cell fate through chromatin reprogramming (Figure 2A; Christensen et al., 2007; Batie et al., 2019). KDM5A also has a relatively low O₂ affinity, with a Km ~90 μM (Table 4; Batie et al., 2019). Inactivation of KDM5A by hypoxia results in rapidly increasing H3K4me3 levels, and downstream effects include active transcriptions of genes associated with antiproliferation, antiapoptosis, etc. (Batie et al., 2019). Excitingly, a new study found that KDM5A binding to H3K4me3 is enhanced by PHD1-mediated hydroxylation of H3 at proline residue 16 (H3P16OH) (Liu et al., 2022). This KDM5A-PHD1 axis raises the possibility of other cross-talk between O₂ sensors.

KDM4 family enzymes mainly catalyze the demethylation at H3K9me2/me3 and H3K36me2 (Table 5; Hillringhaus et al., 2011). Their O₂-binding affinities have also been investigated in vitro: the O₂ Km values of KDM4A, KDM4B, KDM4C, and KDM4E are all within the range of 57–197 μM (Table 4; Cascella and Mirica, 2012; Hancock et al., 2017). These Km values suggest the potential of KDM4 members to be hypoxia sensors, but this depends on their cellular roles and the downstream responses of their speculated inhibition during hypoxia (Figure 2B). Of these KDM4 members, cellular activity of KDM4A is reported to show a graded response to O₂ concentration in U2OS cells and so does the demethylation levels on H3K9me3 (Hancock et al., 2017). It is also reported that KDM4A regulates the transcription of HIF1α through the H3K9 methylation status at HIF1α locus during hypoxia in tumors (Dobrynin et al., 2017). More studies for the function of other KDM4 demethylases during hypoxia are still needed.

There are also cases where the JmjC-containing KDMs function as hypoxia sensors through non-histone substrates. One such example is KDM3A, a histone demethylase for H3K9me2/1 sites (Table 5), whose activity on H3K9me2 is maintained even under severe hypoxia (0.2% O₂), suggesting a high binding affinity with O₂ with this substrate (Yamane et al., 2006; Brauchle et al., 2013; Beyer et al., 2008). However, recently Qian et al. discovered that the demethylation activity of KDM3A on a non-histone substrate, peroxisome proliferator-activated receptor gamma coactivator (PGC-1α) K224me, is inhibited by hypoxia (Figure 2A), with Km ~7.6% O₂ (~75 μM), high enough to function as a hypoxia sensor under physiological conditions (Table 4; Qian et al., 2019). PGC-1α is a transcriptional coactivator that binds with transcriptional factor nuclear respiratory factor (NRF1/2) for activating transcription of nucleus-encoded mitochondrial genes (Qian et al., 2019; Scarpulla et al., 2012). The inhibited activity of KDM3A causes the accumulation of K224 mono-methylation on PCG-1α, which reduces the interaction between PCG-1α and NRF1/2, decreasing mitochondrial biogenesis (Qian et al., 2019). Another example is KDM5C, a histone demethylase for H3K4me2/me3 sites, which can also function as an arginine demethylase (Figure 2A; Li et al., 2022; Iwase et al., 2007). Its demethylation of ULK1 R170me2s site is inhibited by 1% O₂ level in LN229 and several other cell lines (Li et al., 2022). This inhibited demethylation stabilizes ULK1 R170me2s, which further activates ULK1 and induces autophagy as a downstream response (Li et al., 2022). The cases of KDM3A and KDM5C suggest the possibility of other JmjC-containing KDMs to sense and respond to hypoxia through undiscovered non-histone substrates.

Besides KDMs, another set of 2-OG-dependent dioxygenases that act as epigenetic regulators are the TET enzymes (TET1, TET2, and TET3 in humans). These enzymes catalyze the hydroxylation of DNA 5-methylcytosine (5mC) to 5-hydroxymethylcytosine (5hmC) (Ponnaluri et al., 2013; Wu and Zhang, 2017). This facilitates the subsequent demethylation of 5hmC into an unmodified cytosine (Ponnaluri et al., 2013; Wu and Zhang, 2017). Since CpG methylation is typically silencing, TETs tend to promote gene activation. A large variance exists in different reports about the O₂ Km values of TET1 and TET2, ranging from 0.31% and 0.53% (3.0 μM and 5.2 μM) using genomic DNA as substrates to 30 μM using oligonucleotides as substrates (Table 4); however, these measurements show much tighter O₂ binding of TET1 and TET2 compared with the aforementioned reported hypoxia sensors (KDM3A, KDM5A, and KDM6A) (Laukka et al., 2016; Thienpont et al., 2016). Severe hypoxia, such as 0.5% O₂ treatment, is reported to directly impair the cellular activities of TETs, increase DNA hypermethylation, and decrease the expression levels of associated genes (Figure 2A; Thienpont et al., 2016). DNA hypermethylation caused by TETs inhibition also happens during pathophysiological hypoxia found in tumors (Thienpont et al., 2016). Considering their O₂ sensitivity, TETs are more likely to function as hypoxia sensors under extreme hypoxic conditions.

Translational modulators

Translational modulators in the 2-OG-dependent dioxygenases include the mRNA hydroxylases and ribosome hydroxylases.

The most abundant RNA modification is N⁶-methylation of adenosine (m⁶A), which affects the processing, splicing, translation, and degradation of modified mRNAs (Zaccara et al., 2019; Chen et al., 2019). The dynamics of m⁶A modification is coordinated by methyltransferases (so-called ‘writers’), demethylases (‘erasers’), and identifiers (‘readers’) (Zaccara et al., 2019; Chen et al., 2019). Two m⁶A RNA demethylases have been identified thus far: FTO and ALKBH5, with demonstrated demethylase activity in vitro and in vivo, respectively (Zheng et al., 2013; Jia et al., 2011.Zaccara et al., 2019; Zheng et al., 2013; Mauer et al., 2017). Indirect evidence for a sensor role of these enzymes in hypoxia response is that despite their protein levels staying relatively constant, hypoxia leads to m⁶A accumulation in cancer cells and breast cancer. Thus, it is possible that hypoxia plays a role in direct inhibition of ALKBH5 and/or FTO (Figure 2B). More studies about other possible demethylases for m⁶A in mRNA and the O₂-binding affinities of these enzymes are needed to determine which of them directly sense and respond to hypoxia.

The translational apparatus may itself be targeted by ribosome hydroxylases, which modify the histidyl or prolyl residues of ribosomal subunit proteins (Bundred et al., 2018; Zhuang et al., 2015). These ribosome hydroxylases regulate translation and participate in physiological or disease processes, including cellular growth, skeletal bone formation, tumorigenesis, and immune regulation (Bundred et al., 2018; Zhuang et al., 2015; Ge et al., 2012; Singleton et al., 2014). Currently, three ribosome hydroxylases have been identified: histidyl hydroxylases MINA53 (RIOX2) and NO66 (RIOX1) targeting the 60S large subunits Rpl27a and Rpl8, respectively; and prolyl hydroxylase OGFOD1 targeting the 40S small subunit Rpl23 (Ge et al., 2012; Singleton et al., 2014). Among these, NO66 has its activity inhibited by 0.1–1% O₂ in cellular studies (Figure 2B). By contrast, OGFOD1 still retains 80% of its cellular activity even under severe hypoxia (0.2% O₂) (Ge et al., 2012; Singleton et al., 2014). While this suggests the potential for NO66 to be a sensor in severe hypoxia, more studies of the O₂ affinity of NO66 and the downstream response of its inhibition by hypoxia are needed.

Heme-dependent dioxygenases

Heme prosthetic groups are used by these enzymes for O₂ binding and activation (Figure 3—figure supplement 2; Huang and Groves, 2018; Efimov et al., 2011; Raven, 2017). Five known heme-dependent dioxygenases in humans are indoleamine 2,3-dioxygenase (IDO) 1 and 2, tryptophan 2,3-dioxygenase (TDO), and prostaglandin G/H synthase (PGHS) 1 and 2 (Paton and Ntambi, 2009; Efimov et al., 2011; Raven, 2017). Except for IDO2 that has not been measured, these dioxygenases have reported Km values of 10–30 μM (Table 4; Juránek et al., 1999; Kolawole et al., 2015). As reflected by their lower O₂ Km values, heme-dependent dioxygenases tend to have stronger O₂ binding than 2-OG-dependent dioxygenases.

Both TDO and IDOs catalyze the conversion of L-tryptophan to N-formyl-L-kynurenine (Figure 3C; Thackray et al., 2011). They have similar heme- and substrate-binding pockets, although they share low sequence identity overall, and are believed to be an example of convergent evolution (Ball et al., 2014; Thackray et al., 2011). Both TDO and IDO regulate immune responses, possibly by modifying tryptophan homeostasis. TDO and IDO have distinct tissue expression patterns, with TDO mostly restricted to liver and epidermis, while IDO is found throughout the body and can be induced by certain immune or inflammation signals (Ball et al., 2014). Their expression patterns may further regulate their relative importance in different tissues or toward different stimuli (Ball et al., 2014). The cellular activity of TDO is reported to be inhibited by hypoxia (1–10% O₂) in HeLa cells transfected with TDO, while TDO protein level remains unaltered (Elbers et al., 2016). Cellular activity of IDO1 is also decreased by hypoxia (1% O₂) in 86HG39 and HeLa cells with unaltered IDO1 protein level (Schmidt et al., 2013). Impaired immune responses are observed in both cases as downstream effects (Figure 2B; Elbers et al., 2016; Schmidt et al., 2013). Further studies are needed to clarify whether hypoxia directly inhibits the enzymatic activities of TDO and IDO1 and how this might trigger downstream responses in a more physiological system.

Lipoxygenases

Lipoxygenases (LOXs) are iron-containing dioxygenases that catalyze the insertion of O₂ into polyunsaturated fatty acids (PUFA) and their derivatives, forming hydroperoxyl eicosatetraenoic acid (HPETE) products (Figure 3D). HPETE products are chemically unstable and reduced by peroxidases to hydroxyl eicosatetraenoic acid (HETE) (Biringer, 2020; Ivanov et al., 2010; Kuhn et al., 2015). In humans, there are six known LOXs with arachidonic acid as the most common substrate (Biringer, 2020; Kuhn et al., 2015). These arachidonate lipoxygenases (ALOXs) are named according to the positional specificity in their catalyzed hydroperoxyl reactions as the ALOX5, ALOX12, ALOX12B, ALOX15, ALOX15B, and ALOXE3 (Biringer, 2020; Kuhn et al., 2015). Functions of ALOXs include biosynthesis of inflammatory mediators as well as regulation of cellular redox state (Kuhn et al., 2015).

ALOXs demonstrate that O₂-binding affinities can be affected by the specific substrate, as illustrated by ALOX15. The reported O₂ Km values of human ALOX12, rat ALOX5, and rabbit ALOX15 are all within the range of 8–26 μM, as measured by biochemical studies with arachidonic acid as the substrate (Table 4; Juránek et al., 1999; Wecksler et al., 2009). While the rabbit ALOX15 reaches its V_max under normoxia with arachidonic acid as the substrate, its reaction rate with hydroxyl arachidonic acids as substrates still increases with increasing O₂ concentration under hyperoxic conditions (Ivanov et al., 2005). This suggests a higher O₂ Km value, a lower O₂-binding affinity, and the ability to sense the change of O₂ concentration from normoxic to hypoxic conditions when hydroxyl arachidonic acids are the substrates for rabbit ALOX15. Similarly, human ALOX15 has O₂ Km values for different substrates, namely 24 μM for arachidonic acid and 9.6 μM for linoleic acid. Furthermore, allosteric binding of 12-HEHE to ALOX15 affects its O₂ affinity (Wecksler et al., 2009).

The fact that substrates affect O₂ affinity is not a mere laboratory curiosity. Multiple ALOX substrates may be involved in hypoxia-related diseases, including pulmonary hypertension and cardiovascular diseases (Mashima and Okuyama, 2015; Zhu and Ran, 2012; Ivanov et al., 2015). Studying the substrate-dependent O₂ sensitivity of human ALOXs in various biological contexts will be necessary to ascertain whether and how these enzymes sense and respond to hypoxia in vivo.

Other dioxygenases

Ten other human dioxygenases have been identified (Supplementary file 1). They have the shared property of using an octahedral Fe(II) as the catalytic center.

Among these 10 enzymes, cysteamine (2-aminoethanethiol) dioxygenase (ADO) has been identified as a hypoxia sensor (Masson et al., 2019). ADO catalyzes the oxidation of protein N-terminal cysteines to cysteine sulfinic acid (Figure 3E) and promotes the degradation of the oxidized substrate protein through the N-degron pathway (Masson et al., 2019). Human ADO has a relatively low O₂-binding affinity (Km > 500 μM, Table 4, Masson et al., 2019). As a result, even mild hypoxia inhibits ADO activity, allowing stabilization of its substrates, including the regulator of G protein signaling (RGS4/5) and cytokine interleukin (IL)-32 (Masson et al., 2019). During hypoxia, inhibited ADO results in the stabilization of RGS4/5 and subsequently modulates G protein-coupled calcium ion signals and mitogen-activated protein kinase (MAPK) signaling (Figure 2A; Masson et al., 2019). Hypoxia sensing by ADO provides a faster response compared with HIF-mediated transcriptional regulation (Masson et al., 2019).

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