Next Article in Journal
Identification and Monitoring of Amomi Fructus and its Adulterants Based on DNA Barcoding Analysis and Designed DNA Markers
Previous Article in Journal
Synthesis and Antiproliferative Evaluation of Novel Hybrids of Dehydroabietic Acid Bearing 1,2,3-Triazole Moiety
Previous Article in Special Issue
Mechanisms of DNA Replication and Repair: Insights from the Study of G-Quadruplexes
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Molecules
Volume 24
Issue 22
10.3390/molecules24224192
molecules-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Review

ENPP1, an Old Enzyme with New Functions, and Small Molecule Inhibitors—A STING in the Tale of ENPP1

by
Kenneth I. Onyedibe
1,2,
Modi Wang
1 and
Herman O. Sintim
1,2,3,*
1
Chemistry Department, Institute for Drug Discovery, Purdue University, West Lafayette, IN 47907, USA
2
Purdue Institute for Inflammation, Immunology, and Infectious Diseases, West Lafayette, IN 47907, USA
3
Purdue University Center for Cancer Research, West Lafayette, IN 47907, USA
*
Author to whom correspondence should be addressed.
Molecules 2019, 24(22), 4192; https://doi.org/10.3390/molecules24224192
Submission received: 24 September 2019 / Revised: 7 November 2019 / Accepted: 13 November 2019 / Published: 19 November 2019
(This article belongs to the Special Issue Recent Advances in Non-Canonical Nucleic Acid Structures)
Browse Figures
Versions Notes

Abstract

:
Ectonucleotide pyrophosphatase/phosphodiesterase I (ENPP1) was identified several decades ago as a type II transmembrane glycoprotein with nucleotide pyrophosphatase and phosphodiesterase enzymatic activities, critical for purinergic signaling. Recently, ENPP1 has emerged as a critical phosphodiesterase that degrades the stimulator of interferon genes (STING) ligand, cyclic GMP–AMP (cGAMP). cGAMP or analogs thereof have emerged as potent immunostimulatory agents, which have potential applications in immunotherapy. This emerging role of ENPP1 has placed this “old” enzyme at the frontier of immunotherapy. This review highlights the roles played by ENPP1, the mechanism of cGAMP hydrolysis by ENPP1, and small molecule inhibitors of ENPP1 with potential applications in diverse disease states, including cancer.

Graphical Abstract

1. Introduction

Ectonucleotide pyrophosphatase/phosphodiesterase 1 (ENPP1), also called PC-1, is a type II transmembrane glycoprotein with nucleotide pyrophosphatase and phosphodiesterase enzymatic activities [1]. In 1970, Takahashi et al. showed that PC-1 (or ENPP1) was a 115 kDa and 230 kDa protein under reducing and non-reducing conditions, respectively [2]. ENPP1 is expressed in many tissues [3] and is critical for purinergic signaling, which plays an important role in the regulation of cardiovascular, neurological, immunological, musculoskeletal, hormonal, and hematological functions in mammals [4,5].
Specifically, purinergic signaling is involved in platelet aggregation, muscle contraction, cell proliferation, migration, differentiation, and apoptosis, as well as in regulating hypoxia and ischemia in tissues [4]. Purinergic receptors are divided into two major families, P1 and P2 receptors [6]. P1 receptors are mediated by adenosine, while P2 receptors (P2X and P2Y) are mediated by nucleotides, such as ATP and uridine-5’-triphosphate (UTP) [6]. The extracellular levels of these nucleotides are tightly controlled by their hydrolysis, which is mediated by membrane-bound ENPP1 [7]. ENPP1 catalyzes the hydrolysis of ATP or GTP to AMP or GMP, while generating inorganic pyrophosphates (PPi) [8]. Inorganic pyrophosphates inhibit bone and cartilage mineralization [7,9]. Therefore, the generation of PPi by ENPP1 inadvertently makes ENPP1 a central regulator of bone and cartilage development in mammals [7]. Thus, many inherited mineralization, calcium handling or calcification-related disorders have been linked to loss-of-function mutations in ENPP1, including autosomal recessive hypophosphatemic rickets type 2 (ARHR2), ossification of the posterior longitudinal ligament of the spine, generalized arterial calcification of infancy (GACI), arterial calcification due to CD73 deficiency (ACDC), and pseudoxanthoma elasticum (PXE), also referred to as Gronblad-Strandberg Syndrome (GSS) in some reports [10,11,12,13]. Some classical manifestations of these diseases, such as vascular involvement, are common in ACDC, PXE, and GACI [13,14,15]. These conditions are all ectopic mineralization disorders that occur in the presence of ENPP1 mutations, and have led to the hypothesis that PXE and GACI are actually different manifestations of the same clinical spectrum [13,15]. Recently, Staretz-Chacham et al. reported a fatal multisystemic phenotype of GACI that mimics severe congenital infections and was caused by a novel homozygous ENPP1 mutation [16]. Inactivating mutations of ENPP1, which cause ARHR2, have also been shown to increase the transcription of fibroblast growth factor (FGF) 23 in osteoblasts and osteocytes by a mechanism that has not been fully characterized [10,17,18]. Fibroblast growth factor 23 released from osteoblasts and osteocytes regulates vitamin D metabolism and phosphate homeostasis [17,19]. Additionally, Cole diseases, as well as recently characterized forms of dyschromatoses, have been linked to mutations in ENPP1 [11,20]. Interestingly, Chourabi et al. reported that dyschromatoses patients with ENPP1 mutations also consistently had alterations in their melanocyte development and pigmentation signaling pathways [20]. In addition, insulin resistance, a fundamental starting point for most metabolic diseases, has been linked with ENPP1 mutations in several studies [7,21,22,23]. Huesa et al. observed that ENPP1 knockout mice resisted the expected progression to obesity and insulin resistance despite prolonged high-fat diets [24].
Recently, ENPP1 has been found to play an important role in the immunological responses to various stimuli through the cyclic GMP–AMP synthase (cGAS)–stimulator of interferon genes (STING) pathway [25]. Damage associated molecular patterns (DAMPs) as well as pathogen associated molecular patterns (PAMPs) activate the immune system via STING [26]. cGAS senses cytosolic DNA and catalyzes the conversion of GTP and ATP to cyclic GMP–AMP (cGAMP) [27,28]. Subsequently, 2′3′-cGAMP activates STING to initiate an inflammatory response via the TANK-binding kinase 1 (TBK1)–Interferon Regulatory Factor (IRF) 3 pathway to produce type 1 interferons (IFNs) and other cytokines [27]. A link between the cGAS–STING pathway and ENPP1 has emerged whereby the hydrolysis of cGAMP by ENPP1 attenuates cGAS–STING signaling [25,29,30,31,32] (Figure 1a). Li et al. reported that the half-life of cGAMP is largely dependent on ENPP1 by demonstrating a much longer cGAMP half-life in ENPP1 knockout mice [32]. Bisphosphothionate analogs of 2’3’-cGAMP, which are resistant to ENPP1 hydrolysis, were shown to activate STING more than 10-fold, compared to 2′3′-cGAMP [32], thus, implying that delayed or reduced cGAMP hydrolysis by inhibition of ENPP1 would significantly increase the activation of STING. Wang et al. recently showed that ENPP1 inhibition attenuates pseudo-rabies infections through alteration of cGAMP homeostasis [30]. Pseudo-rabies is a viral infection, and it has recently emerged that some other viruses, such as pox virus, possess cytosolic immune nucleases (poxins), which degrade 2’3’-cGAMP and make it unable to activate STING [33]. These poxins are believed to be viral self-protective enzymes that prevent 2′3′-cGAMP from activating STING [33]. Likewise, Dey et al. reported that Mycobacterium tuberculosis (Mtb) could inhibit STING activation and evade host immunity via the concerted action of cyclic dinucleotide phosphodiesterase (CdnP) and ENPP1 (Figure 1b) [31]. Cyclic-di-AMP and cyclic-di-GMP (Figure 2) from invading pathogens also activate the cGAS–STING pathway in the same manner as 2′3′-cGAMP [27].
In addition to being a target for bone [7,34], cardiovascular [35], and metabolic diseases [23], ENPP1 has now emerged as a therapeutic target for cancers, as discussed below [36]. ENPP1 plays a regulatory function in immune cells such as neutrophils, macrophages, dendritic cells, natural killer cells, and B lymphocytes [37,38]. ENPP1 expression is heightened in M2 macrophages in the presence of cancer and promotes tumor growth and spread [30,39]. The location of the gene for ENPP1 is the 6q22-q23 position, which is a region that has been found to be amplified in many tumors, including breast and neural brain cancers [40,41]. Clearly, ENPP1 inhibitors would play an important role in cancer immunotherapy. Although other excellent reviews on ENPP1 inhibitors have been published, these were published before the new role of ENPP1 in modulating the immunomodulatory ligand, cGAMP, was discovered or fully elucidated [42]. Here, we present the current state of knowledge of ENPP1 and inhibitors, which could be used to modulate ENPP1 function.

2. Mechanism of Hydrolysis of ATP and 2′3′-cGAMP by ENPP1

ENPP1 is located on plasma membranes and ER lumen [43]. One of the main substrates of ENPP1 is ATP, a pro-inflammatory signaling molecule. The level of ATP is relatively high in the tumor microenvironment due to the presence of damaged or dying cells [44]. Hydrolysis of ATP by ENPP1 would produce AMP and PPi extracellularly (Figure 3a) to prevent bone mineralization [45,46]. AMP is then dephosphorylated to adenosine by the ecto-5′-nucleotidase (CD73) in the canonical adenosinergic pathway [47].
Kato et al. reported the crystal structures of the ENPP1 extracellular domain bound to four nucleotide monophosphates (MP), (adenosine (AMP), thymidine (TMP), guanosine (GMP), and cytidine (CMP)) at resolutions of 2.7–3.2 Å, in 2012 [21]. The ENPP1 structure has two N-terminal somatomedin B (SMB)-like domains (SMB1 and SMB2), two linker regions (L1 and L2), a catalytic domain, and a nuclease-like domain (Figure 4a). There are three glycosylation sites for ENPP1, which reinforce the domain interaction [21]. It was believed that the SMB-like domains functioned as a transmembrane anchor and were not involved in the enzymatic activity of ENPP1. On the other hand, ENPP1 is a Ca2+- and Zn2+-dependent enzyme, and enzymatic activity is strongly correlated to the concentration of calcium and zinc ions [21]. The nuclease-like domain of ENPP1 contains a calcium ion-binding EF hand motif, which is found in a large family of calcium-binding proteins. The calcium ion is chelated by Asp780, Asp782, Asp784, and Asp788, and the carbonyl group of Arg786 to form an EF hand-like motif (Figure 4b). The catalytic domain chelates two zinc ions in the active site, a location which plays an important role in the function of ENPP1 [21]. The phosphate group of ATP binds in between the two zinc ions to trigger the bond cleavage process and produce AMP and PPi. The nitrogen-6 atom of AMP interacts with Trp304 and Asp308 by a H2O-mediated hydrogen bond network. However, no hydrogen-bonding network is formed for TMP, GMP, and CMP, thereby resulting in the selectivity of ENPP1 for ATP. Hence, ATP is the most efficient and well-investigated substrate for ENPP1. There are other reported natural substrates of ENPP1 including UTP, diadenosine tetraphosphate (AP4A), cyclic adenosine monophosphate (cAMP), and 2′3′-cGAMP, but not 3′3′-cGAMP (Figure 2) [48]. It is already known that cyclic nucleotides, such as cAMP and cGMP, are exported by multidrug resistance proteins (MRPs) [49,50,51]. Recently, it was also shown that some cyclic dinucleotides (such as cyclic-di-AMP) were exported by MRPs [52]. Cyclic nucleotides are degraded by phosphodiesterases (PDEs) 1 to 11 [53]. However, these PDEs do not degrade cyclic dinucleotides, such as 2′3′-cGAMP. Viral poxins and ENPP1 are now the known hydrolytic enzymes of 2′3′-cGAMP [32,33]. ENPP1 hydrolyzes 2′3′-cGAMP with a kinetic rate constant that is similar to the hydrolysis of ATP (Figure 3b) [32].
To understand the mechanism of 2′3′-cGAMP hydrolysis by ENPP1, Kato et al. investigated the crystal structure of ENPP1 in complex with 2′3′-cGAMP (Figure 5) [25]. ENPP1 utilizes two Zn2+ ions, coordinated by Asp358/His362/His517 and Asp200/Asp405/His406, to interact with the phosphate oxygen. The enzyme hydrolyzes the 2′-5′ phosphodiester bond of 2′3′-cGAMP first to form phosphoadenylyl guanosine (pApG) and then a second hydrolysis produces 5′-AMP and 5′-GMP [25]. Recently, Eaglesham et al. described cGAMP hydrolysis by viral and metazoan poxins [33]. However, the mechanism for cGAMP hydrolysis by poxin is different from that of ENPP1. Firstly, cGAMP binds to ENPP1 as the anion and one of the phosphates coordinates to two zinc ions ligated by histidine and aspartate residues, as shown in Figure 5c, whereas, cGAMP binds to poxin as the acid with no active site metal (Figure 6a) [33]. Additionally, for ENPP1, an Oγ atom of a threonine residue (Thr 238) is the nucleophile for hydrolysis, whereas in the poxin mechanism, the nucleophile for phosphate cleavage is the 2’OH of the cGAMP. In addition, for the poxin hydrolysis mechanism, histidine (H17) and tyrosine (Y138) act as general acids, whereas lysine (K142) acts as a general base to deprotonate 2’OH on cGAMP. The first stage of the poxin hydrolysis leads to a 2′-3′-cyclic phosphate, which is then hydrolyzed by an active site hydroxyl ion (OH−), as seen in Figure 6b. Another difference between the poxin and ENPP1 mechanisms of hydrolysis of cGAMP is that ENPP1 cleaves the 2′-5′ phosphodiester bond first, followed by cleavage of the 3′-5′ bonds, whereas poxins only cleave the 3′-5′ phosphodiester bond [25,33]. Notwithstanding these differences, both types of hydrolysis ensure that the hydrolyzed 2′3′-cGAMP is not able to activate the STING pathway.

3. STING and ENPP1 in Cancer

Several studies have shown that STING pathway activation could stimulate anti-inflammatory T cell responses via the induction of indoleamine 2,3 dioxygenase (IDO), which could in turn cause immune suppression in the tumor microenvironment [54,55,56,57,58]. Indoleamine 2,3 dioxygenase 1 (IDO1) is an example of an IFN-stimulated gene (ISG) responsive to IFNs via IFN-response elements found in the mammalian IDO1 gene promoters. IDO is a tryptophan-catabolizing enzyme, which promotes the activation of CD4+ regulatory T cells mainly of the Foxp3-lineage and the subsequent suppression of effector and helper T cell functions [56]. In dendritic cells, IDO enhances immune tolerance by induction of transforming growth factor beta (TGF β) [55,57,59]. In a Lewis lung carcinoma mice model, Lemos et al. showed that knockout of STING caused enhanced killing of cancer cells due to increased CD8+ T cell activity, reduced myeloid suppressor cell infiltration, and high levels of IL10 production in the tumor microenvironment [55]. However, the induction of IDO did not produce the same effect in STING-deficient EL4 thymoma, B16 melanoma, and in neo-antigen-expressing lung carcinoma [55]. The immune-suppressive effect of IDO that leads to tumorigenesis is promoted in tumors with low antigenicity [55]. Hence, when tumor antigenicity is low, STING activation induces immune-regulatory responses via IDO predominantly, whereas, in tumors with high antigenicity, immune-stimulatory responses are enhanced [55]. Similarly, and perhaps, more importantly, adenosine produced by the adenosinergic pathway exhibits significant immunosuppressive effects in the tumor microenvironment and contributes to tumor progression [60,61]. ATP is rapidly dephosphorylated in a stepwise manner in the extracellular milieu by the ecto-nucleotidases, CD39 and CD73 [53]. Typically, CD39 converts ATP to ADP and ADP to AMP, while CD73 dephosphorylates AMP to adenosine (Figure 7) [60,62]. Additionally, in the usually hypoxic tumor microenvironment, hypoxia induces further CD39- and CD73-mediated adenosine production [63,64]. To add to this, hypoxia also inhibits breakdown of adenosine and potentiates adenosine release by downregulating adenosine kinase [61,65]. The increased production of adenosine and the inhibition of its breakdown ultimately leads to much higher levels of adenosine in tumors when compared to normal tissues [66]. The excess adenosine produced by these mechanisms essentially turns off both the innate and adaptive immune responses via G-protein-coupled A2A and A2B adenosine receptors that stimulate cyclic AMP, consequently leading to decreased production of proinflammatory cytokines and increased synthesis of anti-inflammatory cytokines [44,65]. In cervical cancer-derived mesenchymal stromal cells (MSC), de Lourdes Mora-Garcia et al. demonstrated that cytotoxic T lymphocyte effector activity, including proliferation and production of IFN-γ+, were inhibited by adenosine in a dose-dependent manner [65]. Other studies by Garcia-Rocha et al. also reported that MSC derived from cervical cancer tumors induced the expression and secretion of anti-inflammatory cytokines, such as TGF-β1 and IL-10, in cervical cancer cells, thereby protecting the cells from T cell cytotoxicity [67]. The hydrolysis of cGAMP by ENPP1 leads to the production of AMP, which eventually contributes to a more profound immunosuppression via the subsequent dephosphorylation of AMP to adenosine by CD73 [47]. The role of ENPP1 in cancer is exemplified by the observations of enhanced tumor metastasis to the bone from breast cancer, for example, by over-expression of ENPP1 [41]. The significance of ENPP1 and CD73-mediated production of adenosine is further demonstrated by several reports of resistance to carcinogenesis or metastasis by mice deficient in either CD73 or ENPP1 [41,63,68,69].

4. Inhibitors of ENPP1

4.1. Nucleotide-Based Inhibitors of ENPP1

The therapeutic potential of ENPP1 inhibitors has increased with the discovery of ENPP1’s role in modulating the cGAS–STING pathway. In the past years, a few nucleotide-based ENPP1 inhibitors have been developed, which are mostly substrate analogs (Figure 8), such as adenine nucleotide derivatives [71,72,73,74,75]. These adenine nucleotide analog inhibitors of ENPP1 generally exhibit a competitive type of inhibition as their structures are similar to natural ENPP1 substrates [42]. Additionally, the inhibition properties of these nucleotide analogues appear to be similar. Using human soluble ENPP1 and ATP as substrates, Ki for compounds α,β-metADP, α,β-metATP, 2-MeSADP, and 2-MeSATP, bzATP ranged from 13 to 32 µM, and was considered to be moderately potent [42,71,72]. Other nucleotide-based inhibitors have also been reported, such as γ-S-α, β-metATP derivatives, ARL 67156, α-borano-β, γ-metATP derivatives, and diadenosine boranophosphate derivatives. These other nucleotide inhibitors used p-Nph-5′-TMP or p-Nph-5′-AMP as substrates to achieve colorimetric detection of activity [73,74,75].
However, the high acidity precludes oral bioavailability and limits the applicability of nucleotide-based inhibitors. Furthermore, the NPP1 selectivity of these inhibitors against other ectonucleotidases is not well understood. Since the structure of nucleotide-based inhibitors is similar to the structure of natural substrates, the possibility of off-target biological effects, such as P2 purinergic receptor activation, could also be enhanced. Consequently, nucleotide-based inhibitors may not be ideal lead candidates for the development of translatable ENPP1 inhibitors.

4.2. Non-Nucleotide-Based Inhibitors of ENPP1

Many of the reported nucleotide-based inhibitors become negatively charged at physiological pH and have very poor oral bioavailability [76]. Nucleotide-based inhibitors also have very challenging synthesis and purification steps [76]. Non-nucleotide ENPP1 inhibitors have also been reported (Table 1). For example, polyoxometalates [TiW11CoO40]8− were found to be one of the most potent ENPP1 inhibitors, with a Ki of 1.46 nM (0.00146 µM), when compared to other non-nucleotide human soluble enzyme inhibitors, such as reactive blue 2 (RB2), quinazoline derivative, and suramin, with a Ki of 0.141, 0.215, and 0.780 µM, respectively [42]. Among the human membrane-bound enzyme inhibitors with ATP as a substrate, suramin had a superior Ki of 0.26 µM relative to RB2, with a Ki of 0.52 µM. In contrast to nucleotide-based inhibitors, which exhibit competitive inhibition, suramin showed uncompetitive inhibition against the human soluble enzyme [42]. However, heparin, one of the first ENPP1 inhibitors, described about 2 decades ago, was reported to have a relatively high IC50 of 100 µM [77]. Several other non-nucleotide-based inhibitors have been reported, where other substrates, such as ATP with a radioactive phosphorus atom at γ-position of the triphosphate ([γ−32P]ATP), etheno-diadenosine diphosphate, bis(p-nitrophenyl) phosphate, p-nitrophenyl phenyl phosphate, p-Nph-5′-TMP, or p-Nph-5′-AMP were used (Table 1) [42]. The Ki values in these reports varied widely and ranged from 0.0593 to 1400 µM (see Table 1) [71,76,78,79,80,81].
Most recently, in 2019, two new sulfamate derivatives were reported by Semreen et al. and El-Gamal et al., with an IC50 of 0.387 and 0.29 µM, respectively [82,83]. The sulfamate and sulfonate derivate described by El-Gamal was synthesized from a backbone of raloxifene hydrochloride [83]. Raloxifene is an FDA-approved selective estrogen receptor modulator used in the prevention and treatment of postmenopausal osteoporosis, and is known to reduce the risk of breast cancer [84]. Earlier in 2018, Forcellini et al. reported several novel quinazoline-4-piperidine sulfamide analogs as inhibitors of ENPP1 [85]. Among these, meta-pyridine substituted compound 7c was the most potent compound, with a Ki of 58 nM (0.058 µM) [85]. More recently, Weston et al. reported a new ENPP1 inhibitor, SR-8314, which promotes STING activation [86]. The Ki value of SR-8314 against ENPP1 activity was reported as 0.079 µM. It was also shown that SR-8314 has anti-tumor activity with more CD3+, CD4+, and CD8+ T cells found in SR-8314 treatments compared to controls [86]. Baird et al. developed another selective ENPP1 inhibitor, MV-626, which prevents the hydrolysis of cGAMP, and increases STING activation [87]. Therapeutic doses of MV-626 did not show toxicity in mice [87]. In combination with radiation therapy, MV-626 increased overall survival, and the majority of test animals had durable tumor cures [87]. Currently, the ideal doses, intensities, and durations of such therapies has not been established. Generally, hyperactivation of host immune responses and ectopic calcifications by ENPP1 inhibition might be a concern, as discussed in a previous review [36]. However, ENPP1 knockout mice have remained viable, lending credence to the potential use of ENPP1 inhibitors without debilitating adverse outcomes. The addition of these new, more potent inhibitors into the growing array of ENPP1 inhibitors, is a reflection of the therapeutic usefulness of ENPP1 inhibition in immunotherapy.

5. Conclusions

In this review, we have discussed the mechanism and hydrolysis of the relatively new ENPP1 substrate, cGAMP, and that of ATP. We have also highlighted the typical and recently reported ENPP1 inhibitors, based on their classification as either nucleotide-based or non-nucleotide-based inhibitors. Since STING activation is a promising therapeutic strategy to cure cancer, more and more compounds that activate the STING pathway have been reported. ENPP1, as a highly potent cGAMP-degradation enzyme, makes the application of ENPP1 inhibitors for anti-tumor therapy a very topical issue. For virally-infected hosts, small molecule inhibitors of ENPP1 would also need to inhibit viral poxins for maximum efficacy.

Author Contributions

Conceptualization, H.O.S; literature search and methodology, H.O.S., K.I.O. and M.W; writing—original draft preparation, H.O.S, K.I.O. and M.W; writing—review and editing, H.O.S, K.I.O. and M.W.; supervision, H.O.S.; funding acquisition, H.O.S.

Funding

This research was funded by Purdue University.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Evans:, W.H.; Hood, D.O.; Gurd, J.W. Purification and properties of a mouse liver plasma-membrane glycoprotein hydrolysing nucleotide pyrophosphate and phosphodiester bonds. Biochem. J. 1973, 135, 819–826. [Google Scholar] [CrossRef] [PubMed]
  2. Takahashi, T.; Old, L.J.; Boyse, E.A. Surface alloantigens of plasma cells. J. Exp. Med. 1970, 131, 1325–1341. [Google Scholar] [CrossRef] [PubMed]
  3. Huang, R.; Rosenbach, M.; Vaughn, R.; Provvedini, D.; Rebbe, N.; Hickman, S.; Goding, J.; Terkeltaub, R. Expression of the murine plasma cell nucleotide pyrophosphohydrolase PC-1 is shared by human liver, bone, and cartilage cells. Regulation of PC-1 expression in osteosarcoma cells by transforming growth factor-beta. J. Clin. Invest. 1994, 94, 560–567. [Google Scholar] [CrossRef] [PubMed]
  4. Burnstock, G. Purinergic signalling. Br. J. Pharmacol. 2006, 147 (Suppl. 1), S172–S181. [Google Scholar] [CrossRef]
  5. Drury, A.N.; Szent-Györgyi, A. The physiological activity of adenine compounds with especial reference to their action upon the mammalian heart. J. Physiol. 1929, 68, 213–237. [Google Scholar] [CrossRef] [PubMed]
  6. Idzko, M.; Ferrari, D.; Riegel, A.K.; Eltzschig, H.K. Extracellular nucleotide and nucleoside signaling in vascular and blood disease. Blood 2014, 124, 1029–1037. [Google Scholar] [CrossRef]
  7. Roberts, F.; Zhu, D.; Farquharson, C.; Macrae, V.E. ENPP1 in the Regulation of Mineralization and Beyond. Trends Biochem. Sci. 2019, 44, 616–628. [Google Scholar] [CrossRef]
  8. Stefan, C.; Jansen, S.; Bollen, M. NPP-type ectophosphodiesterases: Unity in diversity. Trends Biochem. Sci. 2005, 30, 542–550. [Google Scholar] [CrossRef]
  9. Cimpean, A.; Stefan, C.; Gijsbers, R.; Stalmans, W.; Bollen, M. Substrate-specifying determinants of the nucleotide pyrophosphatases/phosphodiesterases NPP1 and NPP2. Biochem. J. 2004, 381, 71–77. [Google Scholar] [CrossRef]
  10. Lorenz-Depiereux, B.; Schnabel, D.; Tiosano, D.; Häusler, G.; Strom, T.M. Loss-of-function ENPP1 mutations cause both generalized arterial calcification of infancy and autosomal-recessive hypophosphatemic rickets. Am. J. Hum. Genet. 2010, 86, 267–272. [Google Scholar] [CrossRef]
  11. Eytan, O.; Morice-Picard, F.; Sarig, O.; Ezzedine, K.; Isakov, O.; Li, Q.; Ishida-Yamamoto, A.; Shomron, N.; Goldsmith, T.; Fuchs-Telem, D.; et al. Cole Disease Results from Mutations in ENPP1. Am. J. Hum. Genet. 2013, 93, 752–757. [Google Scholar] [CrossRef] [PubMed]
  12. Thumbigere-Math, V.; Alqadi, A.; Chalmers, N.I.; Chavez, M.B.; Chu, E.Y.; Collins, M.T.; Ferreira, C.R.; FitzGerald, K.; Gafni, R.I.; Gahl, W.A.; et al. Hypercementosis Associated with ENPP1 Mutations and GACI. J. Dent. Res. 2018, 97, 432–441. [Google Scholar] [CrossRef] [PubMed]
  13. Li, Q.; van de Wetering, K.; Uitto, J. Pseudoxanthoma Elasticum as a Paradigm of Heritable Ectopic Mineralization Disorders: Pathomechanisms and Treatment Development. Am. J. Pathol. 2019, 189, 216–225. [Google Scholar] [CrossRef] [PubMed]
  14. Van Gils, M.; Nollet, L.; Verly, E.; Deianova, N.; Vanakker, O.M. Cellular signaling in pseudoxanthoma elasticum: An update. Cell. Signal. 2019, 55, 119–129. [Google Scholar] [CrossRef] [PubMed]
  15. Devriese, M.; Legrand, A.; Courtois, M.C.; Jeunemaitre, X.; Albuisson, J. Pseudoxanthoma elasticum with prominent arterial calcifications evoking CD73 deficiency. Vasc. Med. 2019, 24, 461–464. [Google Scholar] [CrossRef] [PubMed]
  16. Staretz-Chacham, O.; Shukrun, R.; Barel, O.; Pode-Shakked, B.; Pleniceanu, O.; Anikster, Y.; Shalva, N.; Ferreira, C.R.; Ben-Haim Kadosh, A.; Richardson, J.; et al. Novel homozygous ENPP1 mutation causes generalized arterial calcifications of infancy, thrombocytopenia, and cardiovascular and central nervous system syndrome. Am. J. Med. Genet. 2019, 179, 2112–2118. [Google Scholar] [CrossRef]
  17. Quarles, L.D. Skeletal secretion of FGF-23 regulates phosphate and vitamin D metabolism. Nat. Rev. Endocrinol. 2012, 8, 276–286. [Google Scholar] [CrossRef]
  18. Levy-Litan, V.; Hershkovitz, E.; Avizov, L.; Leventhal, N.; Bercovich, D.; Chalifa-Caspi, V.; Manor, E.; Buriakovsky, S.; Hadad, Y.; Goding, J.; et al. Autosomal-recessive hypophosphatemic rickets is associated with an inactivation mutation in the ENPP1 gene. Am. J. Hum. Genet. 2010, 86, 273–278. [Google Scholar] [CrossRef]
  19. Quarles, L.D. Evidence for a bone-kidney axis regulating phosphate homeostasis. J. Clin. Invest. 2003, 112, 642–646. [Google Scholar] [CrossRef]
  20. Chourabi, M.; Liew, M.S.; Lim, S.; H’mida-Ben Brahim, D.; Boussofara, L.; Dai, L.; Wong, P.M.; Foo, J.N.; Sriha, B.; Robinson, K.S.; et al. ENPP1 Mutation Causes Recessive Cole Disease by Altering Melanogenesis. J. Invest. Dermatol. 2018, 138, 291–300. [Google Scholar] [CrossRef]
  21. Kato, K.; Nishimasu, H.; Okudaira, S.; Mihara, E.; Ishitani, R.; Takagi, J.; Aoki, J.; Nureki, O. Crystal structure of Enpp1, an extracellular glycoprotein involved in bone mineralization and insulin signaling. Proc. Natl. Acad. Sci. USA 2012, 109, 16876–16881. [Google Scholar] [CrossRef] [PubMed]
  22. Goldfine, I.D.; Maddux, B.A.; Youngren, J.F.; Reaven, G.; Accili, D.; Trischitta, V.; Vigneri, R.; Frittitta, L. The role of membrane glycoprotein plasma cell antigen 1/ectonucleotide pyrophosphatase phosphodiesterase 1 in the pathogenesis of insulin resistance and related abnormalities. Endocr. Rev. 2008, 29, 62–75. [Google Scholar] [CrossRef] [PubMed]
  23. Pan, W.; Ciociola, E.; Saraf, M.; Tumurbaatar, B.; Tuvdendorj, D.; Prasad, S.; Chandalia, M.; Abate, N. Metabolic consequences of ENPP1 overexpression in adipose tissue. Am. J. Physiol. Endocrinol. Metab. 2011, 301, E901–E911. [Google Scholar] [CrossRef] [PubMed]
  24. Huesa, C.; Zhu, D.; Glover, J.D.; Ferron, M.; Karsenty, G.; Milne, E.M.; Millan, J.L.; Ahmed, S.F.; Farquharson, C.; Morton, N.M.; et al. Deficiency of the bone mineralization inhibitor NPP1 protects mice against obesity and diabetes. Dis. Model. Mech. 2014, 7, 1341–1350. [Google Scholar] [CrossRef]
  25. Kato, K.; Nishimasu, H.; Oikawa, D.; Hirano, S.; Hirano, H.; Kasuya, G.; Ishitani, R.; Tokunaga, F.; Nureki, O. Structural insights into cGAMP degradation by Ecto-nucleotide pyrophosphatase phosphodiesterase 1. Nat. Commun. 2018, 9, 4424. [Google Scholar] [CrossRef] [PubMed]
  26. Li, X.D.; Wu, J.; Gao, D.; Wang, H.; Sun, L.; Chen, Z.J. Pivotal roles of cGAS-cGAMP signaling in antiviral defense and immune adjuvant effects. Science 2013, 341, 1390–1394. [Google Scholar] [CrossRef] [PubMed]
  27. Sun, L.; Wu, J.; Du, F.; Chen, X.; Chen, Z.J. Cyclic GMP-AMP synthase is a cytosolic DNA sensor that activates the type I interferon pathway. Science 2013, 339, 786–791. [Google Scholar] [CrossRef]
  28. Wu, J.; Sun, L.; Chen, X.; Du, F.; Shi, H.; Chen, C.; Chen, Z.J. Cyclic GMP-AMP is an endogenous second messenger in innate immune signaling by cytosolic DNA. Science 2013, 339, 826–830. [Google Scholar] [CrossRef]
  29. Haag, S.M.; Gulen, M.F.; Reymond, L.; Gibelin, A.; Abrami, L.; Decout, A.; Heymann, M.; van der Goot, F.G.; Turcatti, G.; Behrendt, R.; et al. Targeting STING with covalent small-molecule inhibitors. Nature 2018, 559, 269–273. [Google Scholar] [CrossRef]
  30. Wang, J.; Lu, S.F.; Wan, B.; Ming, S.L.; Li, G.L.; Su, B.Q.; Liu, J.Y.; Wei, Y.S.; Yang, G.Y.; Chu, B.B. Maintenance of cyclic GMP-AMP homeostasis by ENPP1 is involved in pseudorabies virus infection. Mol. Immunol. 2018, 95, 56–63. [Google Scholar] [CrossRef]
  31. Dey, R.J.; Dey, B.; Zheng, Y.; Cheung, L.S.; Zhou, J.; Sayre, D.; Kumar, P.; Guo, H.; Lamichhane, G.; Sintim, H.O.; et al. Inhibition of innate immune cytosolic surveillance by an M. tuberculosis phosphodiesterase. Nat. Chem. Biol. 2017, 13, 210–217. [Google Scholar] [CrossRef] [PubMed]
  32. Li, L.; Yin, Q.; Kuss, P.; Maliga, Z.; Millán, J.L.; Wu, H.; Mitchison, T.J. Hydrolysis of 2′3′-cGAMP by ENPP1 and design of nonhydrolyzable analogs. Nat. Chem. Biol. 2014, 10, 1043–1048. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  33. Eaglesham, J.B.; Pan, Y.; Kupper, T.S.; Kranzusch, P.J. Viral and metazoan poxins are cGAMP-specific nucleases that restrict cGAS-STING signalling. Nature 2019, 566, 259–263. [Google Scholar] [CrossRef] [PubMed]
  34. Danino, O.; Svetitsky, S.; Kenigsberg, S.; Levin, A.; Journo, S.; Gold, A.; Drexler, M.; Snir, N.; Elkayam, O.; Fischer, B.; et al. Inhibition of nucleotide pyrophosphatase/phosphodiesterase 1: Implications for developing a calcium pyrophosphate deposition disease modifying drug. Rheumatology 2018, 57, 1472–1480. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  35. Nitschke, Y.; Hartmann, S.; Torsello, G.; Horstmann, R.; Seifarth, H.; Weissen--Plenz, G.; Rutsch, F. Expression of NPP1 is regulated during atheromatous plaque calcification. J. Cell. Mol. Med. 2011, 15, 220–231. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  36. Sharma, M.; Thode, T.; Weston, A.; Kaadige, M.R. Development of Enpp1 Inhibitors as a Strategy to Activate Stimulator of Interferon Genes (STING) in Cancers and Other Diseases. Int. J. Cell Sci. Mol. Biol. 2018, 5, 1–5. [Google Scholar]
  37. Abbasi, S.; Shin, D.M.; Beaty, N.; Masiuk, M.; Chen, S.; Gonzalez-Garcia, I.; Zhao, M.; Goding, J.; Morse, H.C.; Wang, H. Characterization of monoclonal antibodies to the plasma cell alloantigen ENPP1. Hybridoma (Larchmt) 2011, 30, 11–17. [Google Scholar] [CrossRef] [Green Version]
  38. Yoon, J.; Wang, H.; Kim, Y.C.; Yoshimoto, M.; Abbasi, S.; Morse Iii, H.C. Plasma cell alloantigen ENPP1 is expressed by a subset of human B cells with potential regulatory functions. Immunol. Cell. Biol. 2016, 94, 719–728. [Google Scholar] [CrossRef]
  39. Kaneda, M.M.; Messer, K.S.; Ralainirina, N.; Li, H.; Leem, C.J.; Gorjestani, S.; Woo, G.; Nguyen, A.V.; Figueiredo, C.C.; Foubert, P.; et al. PI3Kγ is a molecular switch that controls immune suppression. Nature 2016, 539, 437–442. [Google Scholar] [CrossRef] [Green Version]
  40. Grobben, B.; De Deyn, P.P.; Slegers, H. Rat C6 glioma as experimental model system for the study of glioblastoma growth and invasion. Cell Tissue Res. 2002, 310, 257–270. [Google Scholar] [CrossRef]
  41. Lau, W.M.; Doucet, M.; Stadel, R.; Huang, D.; Weber, K.L.; Kominsky, S.L. Enpp1: A potential facilitator of breast cancer bone metastasis. PLoS ONE 2013, 8, e66752. [Google Scholar] [CrossRef] [PubMed]
  42. Lee, S.-Y.; Müller, C.E. Nucleotide pyrophosphatase/phosphodiesterase 1 (NPP1) and its inhibitors. MedChemComm 2017, 8, 823–840. [Google Scholar] [CrossRef] [PubMed]
  43. Bischoff, E.; Tran-Thi, T.-A.; Decker, K.F.A. Nucleotide Pyrophosphatase of Rat Liver. Eur. J. Biochem. 1975, 51, 353–361. [Google Scholar] [CrossRef] [PubMed]
  44. Antonioli, L.; Pacher, P.; Vizi, E.S.; Haskó, G. CD39 and CD73 in immunity and inflammation. Trends. Mol. Med. 2013, 19, 355–367. [Google Scholar] [CrossRef] [Green Version]
  45. Johnson, K.; Goding, J.; Van Etten, D.; Sali, A.; Hu, S.-I.; Farley, D.; Krug, H.; Hessle, L.; Millán, J.L.; Terkeltaub, R. Linked Deficiencies in Extracellular PPi and Osteopontin Mediate Pathologic Calcification Associated With Defective PC-1 and ANK Expression. J. Bone Miner. Res. 2003, 18, 994–1004. [Google Scholar] [CrossRef]
  46. Johnson, K.; Moffa, A.; Chen, Y.; Pritzker, K.; Goding, J.; Terkeltaub, R. Matrix Vesicle Plasma Cell Membrane Glycoprotein-1 Regulates Mineralization by Murine Osteoblastic MC3T3 Cells. J. Bone Miner. Res. 1999, 14, 883–892. [Google Scholar] [CrossRef]
  47. Horenstein, A.L.; Chillemi, A.; Zaccarello, G.; Bruzzone, S.; Quarona, V.; Zito, A.; Serra, S.; Malavasi, F. A CD38/CD203a/CD73 ectoenzymatic pathway independent of CD39 drives a novel adenosinergic loop in human T lymphocytes. OncoImmunology 2013, 2, e26246. [Google Scholar] [CrossRef] [Green Version]
  48. Namasivayam, V.; Lee, S.-Y.; Müller, C.E. The promiscuous ectonucleotidase NPP1: Molecular insights into substrate binding and hydrolysis. Biochim. Biophys. Acta 2017, 1861, 603–614. [Google Scholar] [CrossRef]
  49. Wielinga, P.R.; van der Heijden, I.; Reid, G.; Beijnen, J.H.; Wijnholds, J.; Borst, P. Characterization of the MRP4- and MRP5-mediated Transport of Cyclic Nucleotides from Intact Cells. J. Biol. Chem. 2003, 278, 17664–17671. [Google Scholar] [CrossRef] [Green Version]
  50. Jedlitschky, G.; Burchell, B.; Keppler, D. The Multidrug Resistance Protein 5 Functions as an ATP-dependent Export Pump for Cyclic Nucleotides. J. Biol. Chem. 2000, 275, 30069–30074. [Google Scholar] [CrossRef] [Green Version]
  51. Guo, Y.; Kotova, E.; Chen, Z.-S.; Lee, K.; Hopper-Borge, E.; Belinsky, M.G.; Kruh, G.D. MRP8, ATP-binding Cassette C11 (ABCC11), Is a Cyclic Nucleotide Efflux Pump and a Resistance Factor for Fluoropyrimidines 2′,3′-Dideoxycytidine and 9′-(2′-Phosphonylmethoxyethyl)adenine. J. Biol. Chem. 2003, 278, 29509–29514. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  52. Pham, H.T.; Nhiep, N.T.H.; Vu, T.N.M.; Huynh, T.N.; Zhu, Y.; Huynh, A.L.D.; Chakrabortti, A.; Marcellin, E.; Lo, R.; Howard, C.B.; et al. Enhanced uptake of potassium or glycine betaine or export of cyclic-di-AMP restores osmoresistance in a high cyclic-di-AMP Lactococcus lactis mutant. PLoS Genet. 2018, 14, e1007574. [Google Scholar] [CrossRef] [PubMed]
  53. Conti, M. Phosphodiesterases and cyclic nucleotide signaling in endocrine cells. Mol. Endocrinol. 2000, 14, 1317–1327. [Google Scholar] [CrossRef] [PubMed]
  54. Corrales, L.; McWhirter, S.M.; Dubensky, T.W.; Gajewski, T.F. The host STING pathway at the interface of cancer and immunity. J. Clin. Invest. 2016, 126, 2404–2411. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  55. Lemos, H.; Mohamed, E.; Huang, L.; Ou, R.; Pacholczyk, G.; Arbab, A.S.; Munn, D.; Mellor, A.L. STING Promotes the Growth of Tumors Characterized by Low Antigenicity via IDO Activation. Cancer Res. 2016, 76, 2076–2081. [Google Scholar] [CrossRef] [PubMed]
  56. Lemos, H.; Huang, L.; McGaha, T.L.; Mellor, A.L. Cytosolic DNA sensing via the stimulator of interferon genes adaptor: Yin and Yang of immune responses to DNA. Eur. J. Immunol. 2014, 44, 2847–2853. [Google Scholar] [CrossRef] [Green Version]
  57. Huang, L.; Li, L.; Lemos, H.; Chandler, P.R.; Pacholczyk, G.; Baban, B.; Barber, G.N.; Hayakawa, Y.; McGaha, T.L.; Ravishankar, B.; et al. Cutting edge: DNA sensing via the STING adaptor in myeloid dendritic cells induces potent tolerogenic responses. J. Immunol. 2013, 191, 3509–3513. [Google Scholar] [CrossRef] [Green Version]
  58. Liu, M.; Wang, X.; Wang, L.; Ma, X.; Gong, Z.; Zhang, S.; Li, Y. Targeting the IDO1 pathway in cancer: From bench to bedside. J. Hematol. Oncol. 2018, 11, 100. [Google Scholar] [CrossRef] [Green Version]
  59. Munn, D.H.; Sharma, M.D.; Lee, J.R.; Jhaver, K.G.; Johnson, T.S.; Keskin, D.B.; Marshall, B.; Chandler, P.; Antonia, S.J.; Burgess, R.; et al. Potential regulatory function of human dendritic cells expressing indoleamine 2,3-dioxygenase. Science 2002, 297, 1867–1870. [Google Scholar] [CrossRef]
  60. Vijayan, D.; Young, A.; Teng, M.W.L.; Smyth, M.J. Targeting immunosuppressive adenosine in cancer. Nat. Rev. Cancer 2017, 17, 709–724. [Google Scholar] [CrossRef]
  61. Vigano, S.; Alatzoglou, D.; Irving, M.; Ménétrier-Caux, C.; Caux, C.; Romero, P.; Coukos, G. Targeting Adenosine in Cancer Immunotherapy to Enhance T-Cell Function. Front. Immunol. 2019, 10, 925. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  62. Zimmermann, H.; Zebisch, M.; Sträter, N. Cellular function and molecular structure of ecto-nucleotidases. Purinergic Signal. 2012, 8, 437–502. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  63. Allard, B.; Longhi, M.S.; Robson, S.C.; Stagg, J. The ectonucleotidases CD39 and CD73: Novel checkpoint inhibitor targets. Immunol. Rev. 2017, 276, 121–144. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  64. Rao, S.R.; Snaith, A.E.; Marino, D.; Cheng, X.; Lwin, S.T.; Orriss, I.R.; Hamdy, F.C.; Edwards, C.M. Tumour-derived alkaline phosphatase regulates tumour growth, epithelial plasticity and disease-free survival in metastatic prostate cancer. Br. J. Cancer 2017, 116, 227–236. [Google Scholar] [CrossRef]
  65. de Lourdes Mora-García, M.; García-Rocha, R.; Morales-Ramírez, O.; Montesinos, J.J.; Weiss-Steider, B.; Hernández-Montes, J.; Ávila-Ibarra, L.R.; Don-López, C.A.; Velasco-Velázquez, M.A.; Gutiérrez-Serrano, V.; et al. Mesenchymal stromal cells derived from cervical cancer produce high amounts of adenosine to suppress cytotoxic T lymphocyte functions. J. Transl. Med. 2016, 14, 302. [Google Scholar] [CrossRef] [Green Version]
  66. Montalbán Del Barrio, I.; Penski, C.; Schlahsa, L.; Stein, R.G.; Diessner, J.; Wöckel, A.; Dietl, J.; Lutz, M.B.; Mittelbronn, M.; Wischhusen, J.; et al. Adenosine-generating ovarian cancer cells attract myeloid cells which differentiate into adenosine-generating tumor associated macrophages—A self-amplifying, CD39- and CD73-dependent mechanism for tumor immune escape. J. Immunother. Cancer 2016, 4, 49. [Google Scholar] [CrossRef] [Green Version]
  67. García-Rocha, R.; Moreno-Lafont, M.; Mora-García, M.L.; Weiss-Steider, B.; Montesinos, J.J.; Piña-Sánchez, P.; Monroy-García, A. Mesenchymal stromal cells derived from cervical cancer tumors induce TGF-β1 expression and IL-10 expression and secretion in the cervical cancer cells, resulting in protection from cytotoxic T cell activity. Cytokine 2015, 76, 382–390. [Google Scholar] [CrossRef]
  68. Stagg, J.; Beavis, P.A.; Divisekera, U.; Liu, M.C.; Möller, A.; Darcy, P.K.; Smyth, M.J. CD73-deficient mice are resistant to carcinogenesis. Cancer Res. 2012, 72, 2190–2196. [Google Scholar] [CrossRef]
  69. Stagg, J.; Divisekera, U.; Duret, H.; Sparwasser, T.; Teng, M.W.; Darcy, P.K.; Smyth, M.J. CD73-deficient mice have increased antitumor immunity and are resistant to experimental metastasis. Cancer Res. 2011, 71, 2892–2900. [Google Scholar] [CrossRef] [Green Version]
  70. Lapa, F.D.; Júnior, S.J.; Cerutti, M.L.; Santos, A.R. Pharmacology and Therapeutics. In Pharmacology of Adenosine Receptors and Their Signaling Role in Immunity and Inflammation; Gowder, S.J.T., Ed.; IntechOpen: London, UK, 2014; pp. 86–130. [Google Scholar]
  71. Lee, S.-Y.; Sarkar, S.; Bhattarai, S.; Namasivayam, V.; De Jonghe, S.; Stephan, H.; Herdewijn, P.; El-Tayeb, A.; Müller, C.E. Substrate-Dependence of Competitive Nucleotide Pyrophosphatase/Phosphodiesterase1 (NPP1) Inhibitors. Front. Pharmacol. 2017, 8. [Google Scholar] [CrossRef] [Green Version]
  72. Lee, S.-Y.; Müller, C.E. Large-volume sample stacking with polarity switching for monitoring of nucleotide pyrophosphatase/phosphodiesterase 1 (NPP1) reactions by capillary electrophoresis. Electrophoresis 2014, 35, 855–863. [Google Scholar] [CrossRef] [PubMed]
  73. Nadel, Y.; Lecka, J.; Gilad, Y.; Ben-David, G.; Förster, D.; Reiser, G.; Kenigsberg, S.; Camden, J.; Weisman, G.A.; Senderowitz, H.; et al. Highly Potent and Selective Ectonucleotide Pyrophosphatase/Phosphodiesterase I Inhibitors Based on an Adenosine 5′-(α or γ)-Thio-(α,β- or β,γ)-methylenetriphosphate Scaffold. J. Med. Chem. 2014, 57, 4677–4691. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  74. Lévesque, S.A.; Lavoie, É.G.; Lecka, J.; Bigonnesse, F.; Sévigny, J. Specificity of the ecto-ATPase inhibitor ARL 67156 on human and mouse ectonucleotidases. Br. J. Pharmacol. 2007, 152, 141–150. [Google Scholar] [CrossRef] [PubMed]
  75. Lecka, J.; Ben-David, G.; Simhaev, L.; Eliahu, S.; Oscar, J.; Luyindula, P.; Pelletier, J.; Fischer, B.; Senderowitz, H.; Sévigny, J. Nonhydrolyzable ATP Analogues as Selective Inhibitors of Human NPP1: A Combined Computational/Experimental Study. J. Med. Chem. 2013, 56, 8308–8320. [Google Scholar] [CrossRef] [PubMed]
  76. Chang, L.; Lee, S.Y.; Leonczak, P.; Rozenski, J.; De Jonghe, S.; Hanck, T.; Müller, C.E.; Herdewijn, P. Imidazopyridine- and purine-thioacetamide derivatives: Potent inhibitors of nucleotide pyrophosphatase/phosphodiesterase 1 (NPP1). J. Med. Chem. 2014, 57, 10080–10100. [Google Scholar] [CrossRef] [PubMed]
  77. Hosoda, N.; Hoshino, S.I.; Kanda, Y.; Katada, T. Inhibition of phosphodiesterase/pyrophosphatase activity of PC-1 by its association with glycosaminoglycans. Eur. J. Biochem. 1999, 265, 763–770. [Google Scholar] [CrossRef] [Green Version]
  78. Lee, S.Y.; Perotti, A.; De Jonghe, S.; Herdewijn, P.; Hanck, T.; Müller, C.E. Thiazolo[3,2-a]benzimidazol-3(2H)-one derivatives: Structure-activity relationships of selective nucleotide pyrophosphatase/phosphodiesterase1 (NPP1) inhibitors. Bioorg. Med. Chem. 2016, 24, 3157–3165. [Google Scholar] [CrossRef]
  79. Grobben, B.; Claes, P.; Roymans, D.; Esmans, E.L.; Van Onckelen, H.; Slegers, H. Ecto-nucleotide pyrophosphatase modulates the purinoceptor-mediated signal transduction and is inhibited by purinoceptor antagonists. Br. J. Pharmacol. 2000, 130, 139–145. [Google Scholar] [CrossRef] [Green Version]
  80. Shayhidin, E.E.; Forcellini, E.; Boulanger, M.C.; Mahmut, A.; Dautrey, S.; Barbeau, X.; Lagüe, P.; Sévigny, J.; Paquin, J.F.; Mathieu, P. Quinazoline-4-piperidine sulfamides are specific inhibitors of human NPP1 and prevent pathological mineralization of valve interstitial cells. Br. J. Pharmacol. 2015, 172, 4189–4199. [Google Scholar] [CrossRef] [Green Version]
  81. Ausekle, E.; Ejaz, S.A.; Khan, S.U.; Ehlers, P.; Villinger, A.; Lecka, J.; Sévigny, J.; Iqbal, J.; Langer, P. New one-pot synthesis of N-fused isoquinoline derivatives by palladium-catalyzed C-H arylation: Potent inhibitors of nucleotide pyrophosphatase-1 and -3. Org. Biomol. Chem. 2016, 14, 11402–11414. [Google Scholar] [CrossRef]
  82. Semreen, M.H.; El-Gamal, M.I.; Ullah, S.; Jalil, S.; Zaib, S.; Anbar, H.S.; Lecka, J.; Sévigny, J.; Iqbal, J. Synthesis, biological evaluation, and molecular docking study of sulfonate derivatives as nucleotide pyrophosphatase/phosphodiesterase (NPP) inhibitors. Bioorg. Med. Chem. 2019, 27, 2741–2752. [Google Scholar] [CrossRef] [PubMed]
  83. El-Gamal, M.I.; Ullah, S.; Zaraei, S.O.; Jalil, S.; Zaib, S.; Zaher, D.M.; Omar, H.A.; Anbar, H.S.; Pelletier, J.; Sévigny, J.; et al. Synthesis, biological evaluation, and docking studies of new raloxifene sulfonate or sulfamate derivatives as inhibitors of nucleotide pyrophosphatase/phosphodiesterase. Eur. J. Med. Chem. 2019, 181, 111560. [Google Scholar] [CrossRef] [PubMed]
  84. Rey, J.R.; Cervino, E.V.; Rentero, M.L.; Crespo, E.C.; Alvaro, A.O.; Casillas, M. Raloxifene: Mechanism of action, effects on bone tissue, and applicability in clinical traumatology practice. Open Orthop. J. 2009, 3, 14–21. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  85. Forcellini, E.; Boutin, S.; Lefebvre, C.-A.; Shayhidin, E.E.; Boulanger, M.-C.; Rhéaume, G.; Barbeau, X.; Lagüe, P.; Mathieu, P.; Paquin, J.-F. Synthesis and biological evaluation of novel quinazoline-4-piperidinesulfamide derivatives as inhibitors of NPP1. Eur. J. Med. Chem. 2018, 147, 130–149. [Google Scholar] [CrossRef]
  86. Weston, A.; Thode, T.; Munoz, R.; Daniel, S.; Soldi, R.; Kaadige, M.; Han, H.; Vankayalapti, H.; Sharma, S. Abstract 3077: Preclinical studies of SR-8314, a highly selective ENPP1 inhibitor and an activator of STING pathway. Cancer Res. 2019, 79, 3077. [Google Scholar] [CrossRef]
  87. Baird, J.; Dietsch, G.; Florio, V.; Gallatin, M.; Knox, C.; Odingo, J.; Crittenden, M.; Gough, M.J. MV-626, a potent and selective inhibitor of ENPP1 enhances STING activation and augments T-cell mediated anti-tumor activity in vivo. In Proceedings of the 33rd Annual Meeting of the Society for Immunotherapy of Cancer, Walter E. Washington Convention Center, Washington, DC, USA, 7–11 November 2018; p. 410. [Google Scholar]
  88. Asensio, A.C.; Rodríguez-Ferrer, C.R.; Castañeyra-Perdomo, A.; Oaknin, S.; Rotllán, P. Biochemical analysis of ecto-nucleotide pyrophosphatase phosphodiesterase activity in brain membranes indicates involvement of NPP1 isoenzyme in extracellular hydrolysis of diadenosine polyphosphates in central nervous system. Neurochem. Int. 2007, 50, 581–590. [Google Scholar] [CrossRef]
  89. Iqbal, J.; Lévesque, S.A.; Sévigny, J.; Müller, C.E. A highly sensitive CE-UV method with dynamic coating of silica-fused capillaries for monitoring of nucleotide pyrophosphatase/phosphodiesterase reactions. Electrophoresis 2008, 29, 3685–3693. [Google Scholar] [CrossRef]
  90. Jafari, B.; Yelibayeva, N.; Ospanov, M.; Ejaz, S.A.; Afzal, S.; Khan, S.U.; Abilov, Z.A.; Turmukhanova, M.Z.; Kalugin, S.N.; Safarov, S.; et al. Synthesis of 2-arylated thiadiazolopyrimidones by Suzuki–Miyaura cross-coupling: A new class of nucleotide pyrophosphatase (NPPs) inhibitors. RSC Adv. 2016, 6, 107556–107571. [Google Scholar] [CrossRef]
  91. Lee, S.Y.; Fiene, A.; Li, W.; Hanck, T.; Brylev, K.A.; Fedorov, V.E.; Lecka, J.; Haider, A.; Pietzsch, H.J.; Zimmermann, H.; et al. Polyoxometalates—Potent and selective ecto-nucleotidase inhibitors. Biochem. Pharmacol. 2015, 93, 171–181. [Google Scholar] [CrossRef]
  92. Choudhary, M.I.; Fatima, N.; Khan, K.M.; Jalil, S.; Iqbal, S.; Atta-Ur-Rahman. New biscoumarin derivatives-cytotoxicity and enzyme inhibitory activities. Bioorg. Med. Chem. 2006, 14, 8066–8072. [Google Scholar] [CrossRef]
  93. Khan, K.M.; Fatima, N.; Rasheed, M.; Jalil, S.; Ambreen, N.; Perveen, S.; Choudhary, M.I. 1,3,4-Oxadiazole-2(3H)-thione and its analogues: A new class of non-competitive nucleotide pyrophosphatases/phosphodiesterases 1 inhibitors. Bioorg. Med. Chem. 2009, 17, 7816–7822. [Google Scholar] [CrossRef] [PubMed]
  94. Raza, R.; Akhtar, T.; Hameed, S.; Lecka, J.; Iqbal, J.; Sévigny, J. Identification of Potent and Selective Human Ecto-Nucleotide Pyrophosphatase/Phosphodiesterase-3 (hNPP3) Inhibitors. Open Enzym. Inhib. J. 2011, 4, 17–22. [Google Scholar]
  95. Patel, S.D.; Habeski, W.M.; Cheng, A.C.; de la Cruz, E.; Loh, C.; Kablaoui, N.M. Quinazolin-4-piperidin-4-methyl sulfamide PC-1 inhibitors: Alleviating hERG interactions through structure based design. Bioorg. Med. Chem. Lett. 2009, 19, 3339–3343. [Google Scholar] [CrossRef] [PubMed]
  96. Khan, K.M.; Siddiqui, S.; Saleem, M.; Taha, M.; Saad, S.M.; Perveen, S.; Choudhary, M.I. Synthesis of triazole Schiff bases: Novel inhibitors of nucleotide pyrophosphatase/phosphodiesterase-1. Bioorg. Med. Chem. 2014, 22, 6509–6514. [Google Scholar] [CrossRef]
Molecules 24 04192 g001 550
Figure 1. (a) Scheme of Ectonucleotide pyrophosphatase/phosphodiesterase 1 (ENPP1) function in the cyclic GMP–AMP synthase (cGAS)–stimulator of interferon genes (STING) pathway. (b) Scheme for cyclic dinucleotide signaling and inhibition of cGAS–STING pathway by M. tuberculosis’ cyclic dinucleotide phosphodiesterase (CdnP). Figure 1b adapted from Reference [24] with permission from Springer Nature Limited, Copyright 2016.
Figure 1. (a) Scheme of Ectonucleotide pyrophosphatase/phosphodiesterase 1 (ENPP1) function in the cyclic GMP–AMP synthase (cGAS)–stimulator of interferon genes (STING) pathway. (b) Scheme for cyclic dinucleotide signaling and inhibition of cGAS–STING pathway by M. tuberculosis’ cyclic dinucleotide phosphodiesterase (CdnP). Figure 1b adapted from Reference [24] with permission from Springer Nature Limited, Copyright 2016.
Molecules 24 04192 g001
Molecules 24 04192 g002 550
Figure 2. Structures of different cyclic dinucleotides: c-di-GMP, c-di-AMP, 2′3′-cGAMP, and 3′3′-cGAMP.
Figure 2. Structures of different cyclic dinucleotides: c-di-GMP, c-di-AMP, 2′3′-cGAMP, and 3′3′-cGAMP.
Molecules 24 04192 g002
Molecules 24 04192 g003 550
Figure 3. Hydrolysis of substrates (a) ATP and (b) 2′3′-cGAMP by ENPP1.
Figure 3. Hydrolysis of substrates (a) ATP and (b) 2′3′-cGAMP by ENPP1.
Molecules 24 04192 g003
Molecules 24 04192 g004 550
Figure 4. (a) Domain organization of mouse ENPP1. (b) Crystal structure of the extracellular domain of ENPP1 in complex with AMP. Catalytic domain, cyan; nuclease-like domain, magenta; L1, wheat; L2, yellow-green; EF hand-like motif, pink; insertion loop, gold. AMP and N-glycans are shown as green and yellow sticks, respectively. The bound zinc and calcium ions are shown as gray and yellow-green spheres, respectively. Disulfide linkages are shown as sticks. The two somatomedin B (SMB)-like domains are shown by circles, as they are disordered in the crystal structure [21]. Reproduced from Reference [21] with permission from National Academy of Sciences, Copyright 2012.
Figure 4. (a) Domain organization of mouse ENPP1. (b) Crystal structure of the extracellular domain of ENPP1 in complex with AMP. Catalytic domain, cyan; nuclease-like domain, magenta; L1, wheat; L2, yellow-green; EF hand-like motif, pink; insertion loop, gold. AMP and N-glycans are shown as green and yellow sticks, respectively. The bound zinc and calcium ions are shown as gray and yellow-green spheres, respectively. Disulfide linkages are shown as sticks. The two somatomedin B (SMB)-like domains are shown by circles, as they are disordered in the crystal structure [21]. Reproduced from Reference [21] with permission from National Academy of Sciences, Copyright 2012.
Molecules 24 04192 g004
Molecules 24 04192 g005 550
Figure 5. (a) Overall structure of ENPP1 in complex with phosphoadenylyl guanosine (pApG). The N-linked sugars are shown as yellow sticks. (b) Binding of pApG to the ENPP1 active site. (c) Proposed mechanism of the ENPP1-catalyzed 2′3′-cGAMP degradation [25]. Reproduced from Reference [25] with permission from Springer Nature Limited, Copyright 2018.
Figure 5. (a) Overall structure of ENPP1 in complex with phosphoadenylyl guanosine (pApG). The N-linked sugars are shown as yellow sticks. (b) Binding of pApG to the ENPP1 active site. (c) Proposed mechanism of the ENPP1-catalyzed 2′3′-cGAMP degradation [25]. Reproduced from Reference [25] with permission from Springer Nature Limited, Copyright 2018.
Molecules 24 04192 g005
Molecules 24 04192 g006 550
Figure 6. (a) Crystal structure of Vaccinia virus poxin–2′,3′-cGAMP complex highlighting residues involved in 3′-5′-bond hydrolysis. (b) Mechanism of poxin metal-independent hydrolysis [33]. Reproduced from Reference [33] with permission from Springer Nature Limited, Copyright 2019.
Figure 6. (a) Crystal structure of Vaccinia virus poxin–2′,3′-cGAMP complex highlighting residues involved in 3′-5′-bond hydrolysis. (b) Mechanism of poxin metal-independent hydrolysis [33]. Reproduced from Reference [33] with permission from Springer Nature Limited, Copyright 2019.
Molecules 24 04192 g006
Molecules 24 04192 g007 550
Figure 7. Scheme of the purinergic system in inflammation. Intracellular adenosine is produced from AMP or S-adenosylhomocysteine, which is then exported to the extracellular environment by nucleoside transporters, to stimulate several adenosine receptor-mediated signaling pathways. Extracellular adenosine acts as the signaling molecule that binds and stimulates the cell surface adenosine receptors (A1, A2A, A2B, and A3) and triggers both pro-inflammatory and anti-inflammatory responses. A1, adenosine A1 receptor; A2A, adenosine A2A receptor; A2B, adenosine A2B receptor; A3, adenosine A3 receptor; AC, adenylate cyclase; cAMP, cyclic adenosine monophosphate; ERK1/2, extracellular signal-regulated kinases 1 and 2; IP3, inositol triphosphate; MAPK, mitogen-activated protein kinase; PKB, protein kinase B; PLC, phospholipase C; PI3K, Phosphatidylinositide 3-kinase; ATP, adenosine triphosphate; AMP, adenosine monophosphate; HXT, hypoxanthine; AK, adenosine kinase; ADA, adenosine deaminase; PNP, purine nucleoside phosphorylase; ENTs, equilibrate nucleoside transporters; CD39, ectonucleoside triphosphate diphosphohydrolase; Inhibition →; Activation - - →[70]. Reproduced from Reference [70] with permission from Intech Open, Copyright 2014.
Figure 7. Scheme of the purinergic system in inflammation. Intracellular adenosine is produced from AMP or S-adenosylhomocysteine, which is then exported to the extracellular environment by nucleoside transporters, to stimulate several adenosine receptor-mediated signaling pathways. Extracellular adenosine acts as the signaling molecule that binds and stimulates the cell surface adenosine receptors (A1, A2A, A2B, and A3) and triggers both pro-inflammatory and anti-inflammatory responses. A1, adenosine A1 receptor; A2A, adenosine A2A receptor; A2B, adenosine A2B receptor; A3, adenosine A3 receptor; AC, adenylate cyclase; cAMP, cyclic adenosine monophosphate; ERK1/2, extracellular signal-regulated kinases 1 and 2; IP3, inositol triphosphate; MAPK, mitogen-activated protein kinase; PKB, protein kinase B; PLC, phospholipase C; PI3K, Phosphatidylinositide 3-kinase; ATP, adenosine triphosphate; AMP, adenosine monophosphate; HXT, hypoxanthine; AK, adenosine kinase; ADA, adenosine deaminase; PNP, purine nucleoside phosphorylase; ENTs, equilibrate nucleoside transporters; CD39, ectonucleoside triphosphate diphosphohydrolase; Inhibition →; Activation - - →[70]. Reproduced from Reference [70] with permission from Intech Open, Copyright 2014.
Molecules 24 04192 g007
Molecules 24 04192 g008 550
Figure 8. Structures of nucleotide-based ENPP1 inhibitors.
Figure 8. Structures of nucleotide-based ENPP1 inhibitors.
Molecules 24 04192 g008
Table
Table 1. Non-nucleotide-based inhibitors ‡.
Table 1. Non-nucleotide-based inhibitors ‡.
Inhibitor (I)StructureSubstrateEnzymeKi/ IC50aInhibition Ref
(µM)Type
Polysaccharides Molecules 24 04192 i001
HeparinATPHuman, soluble100aCompetitive [77]
p-Nph-5′-TMPHuman, soluble1.0aCompetitive[77]
εAP2AUnspecified rat NPPs25Competitive[88]
Polysulfonates
Suramin Molecules 24 04192 i002P32-ATPUnspecified rat NPPs61–83aNot defined[79]
ATPHuman, membrane-bound0.26Not defined[89]
p-Nph-5′-TMPHuman, membrane-bound8.67aNot defined[90]
p-Nph-5′-TMPHuman, soluble1.07Uncompetitive[71]
p-Nph-5′-AMP Human, soluble1.03Uncompetitive[71]
ATPHuman, soluble0.78Uncompetitive[71]
Reactive blue 2 Molecules 24 04192 i003P32-ATPUnspecified rat NPPs9–15aNot defined[79]
ATPHuman, membrane-bound0.52Not defined[89]
ATPHuman, soluble0.14Noncompetitive[71]
p-Nph-5′-TMPHuman, soluble0.198Noncompetitive[71]
p-Nph-5′-AMPHuman, soluble0.176Noncompetitive[71]
Polyoxometalates
[TiW11CoO40]8− ATPHuman, soluble0.00146Noncompetitive[91]
Other Heterocyclic compounds
PPADS Molecules 24 04192 i004P32-ATPUnspecified rat NPPs9–15aNot defined[79]
Biscoumarin derivative Molecules 24 04192 i005p-NPPPHuman, soluble50–1000Noncompetitive[92]
bis(p-NP)PSnake venom8–1150Noncompetitive[92]
Oxadiazole derivatives Molecules 24 04192 i006p-NPPPHuman, soluble150–850Noncompetitive[93]
bis(p-NP)PSnake venom100–1400Noncompetitive[93]
p-Nph-5′-TMPHuman, membrane-bound1.9–5.5aNoncompetitive[94]
Quinazoline derivative Molecules 24 04192 i007ATPNot determined0.036–5.98aNot defined[95]
p-Nph-5′-TMPHuman, membrane-bound0.059–0.110Noncompetitive[80]
p-Nph-5′-TMPHuman, soluble0.06bCompetitive[71]
p-Nph-5′-AMPHuman, soluble0.42bCompetitive[71]
ATPHuman, soluble0.215Competitive[71]
Triazole derivative Molecules 24 04192 i008bis(p-NP)PSnake venom132–1164aNot defined[96]
Thioacetamide derivative Molecules 24 04192 i009ATPHuman, soluble5.34–89.7Competitive[76]
p-Nph-5′-TMPHuman, soluble0.005–11.0Competitive[76]
Isoquinoline derivative Molecules 24 04192 i010p-Nph-5′-AMPHuman, soluble14.9Competitive[71]
Thiadiazolopyrimidinone derivative Molecules 24 04192 i011p-Nph-5′-TMPHuman, membrane-bound0.36–2.81aCompetitive[81]
p-Nph-5′-TMPHuman, membrane-bound0.31–2.26aCompetitive[90]
Thiazolobenzimidazolon-e derivative Molecules 24 04192 i012ATPHuman, soluble0.467–0.981 Uncompetitive[78]
Sulfamate derivatives Molecules 24 04192 i013p-Nph-5′-TMPHuman, soluble0.29a [83]
p-Nph-5′-TMP 0.387aCompetitive[82]
SR 8314Not disclosedATPHuman, soluble0.079Not determined[86]
MV 626Not disclosedATP 5–18 [87]
‡ Adapted from Lee et al. 2017 [42] (updated with more recent inhibitors) with permission from the Royal Society of Chemistry, Copyright 2017; a IC50 values, b Value determined with the most potent derivative of quinazolines.

Share and Cite

MDPI and ACS Style

Onyedibe, K.I.; Wang, M.; Sintim, H.O. ENPP1, an Old Enzyme with New Functions, and Small Molecule Inhibitors—A STING in the Tale of ENPP1. Molecules 2019, 24, 4192. https://doi.org/10.3390/molecules24224192

AMA Style

Onyedibe KI, Wang M, Sintim HO. ENPP1, an Old Enzyme with New Functions, and Small Molecule Inhibitors—A STING in the Tale of ENPP1. Molecules. 2019; 24(22):4192. https://doi.org/10.3390/molecules24224192

Chicago/Turabian Style

Onyedibe, Kenneth I., Modi Wang, and Herman O. Sintim. 2019. "ENPP1, an Old Enzyme with New Functions, and Small Molecule Inhibitors—A STING in the Tale of ENPP1" Molecules 24, no. 22: 4192. https://doi.org/10.3390/molecules24224192

Article Metrics

Back to TopTop