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. Author manuscript; available in PMC: 2013 Nov 18.
Published in final edited form as: Methods Enzymol. 2012;516:10.1016/B978-0-12-394291-3.00031-9. doi: 10.1016/B978-0-12-394291-3.00031-9

Fe(II)-dependent, uridine-5′-monophosphate α-ketoglutarate dioxygenases in the synthesis of 5′-modified nucleosides

Zhaoyong Yang 1, Jason Unrine 2, Koichi Nonaka 3, Steven G Van Lanen 4,*
PMCID: PMC3831618  NIHMSID: NIHMS525022  PMID: 23034228

Abstract

Several nucleoside antibiotics from various actinomycetes contain a high-carbon sugar nucleoside that is putatively derived via C-5′-modification of the canonical nucleoside. Two prominent examples are the 5′-C-carbamoyluridine-containing and 5′-C-glycyluridine-containing nucleosides, both families of which were discovered using screens aimed at finding inhibitors of bacterial translocase I involved in the assembly of the bacterial peptidoglycan cell wall. A shared open reading frame was identified whose gene product is similar to enzymes of the non-haem, Fe(II)- and a-ketoglutarate-dependent dioxygenases. The enzyme LipL from the biosynthetic pathway for A-90289, a 5′-C-glycyluridine-containing nucleoside, was functionally characterized as UMP:α-ketoglutarate dioxygenase, providing the enzymatic imperative for the generation of a nucleoside-5′-aldehdye that serves as a downstream substrate for an aldol or aldol-type reaction leading to the high-carbon sugar scaffold. The functional assignment of LipL and the homologous enzymes—including bioinformatic analysis, iron detection and quantification, and assay development for biochemical characterization—is presented herein.

Keywords: Nucleoside, antibiotic, dioxygenase, phosphate, uridine-5′-monophosphate, iron, α-ketoglutarate

1. Introduction

More than 200, structurally unique nucleoside antibiotics of microbial origin have been described with diverse biological activity, including antibacterial, antitumor, and antiviral, among others (Isono, 1991). Although often simple, a significant number of these molecules contain both highly-modified and unusually modified moieties, suggesting that several intriguing biochemical transformations occur during their biosynthesis. One such family is the high-carbon sugar nucleosides, which are modified at C-5′ of the parent ribose to generate furanosides containing 6–11 contiguous carbons. This family includes: C6-furanosyl nucleosides such as capuramycins A-500359s (Muramatsu et al., 2003) and A-503083s (Muramatsu et al., 2004), polyoxins (Isono et al., 1965), and nikkomycins (Hagenmaier et al., 1979); C7-furanosyl nucleosides such as A-90289s (Fujita et al., 2011), caprazamycins (Igarashi et al., 2005) liposidomycins (Ubukata et al., 1992), muraymycins (McDonald et al., 2002), and FR-900453 (Ochi et al., 1989); C8-furanosyl nucleosides such as griseolic acids (Takahashi et al., 1985), octosyl acids (Isono et al., 1975), and ezomycins (Sakata et al., 1975); the C10-furanosyl nucleoside sinefungin (Boeck et al., 1973); and the C11-furansoyl nucleosides herbicidins (Haneishi et al., 1976) and tunicamycins (Takatsuki et al., 1971) (representative structures are shown in Figure 1).

Figure 1. Structures of high-carbon sugar nucleosides.

Figure 1

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Representative structures for (A) C6-, (B) C7-, (C) C8-, (D) C10-, and (E) C11-furanosyl nucleoside antibiotics.

Isotopic enrichment studies using different high-carbon sugar nucleosides as biosynthetic models suggested that the glycosidic bond is established prior to C-5′ modification. In other words, a nucleoside scaffold is the direct precursor that is utilized in a C-C bond-forming event with a separate precursor that—in some instances—is potentially a C2-unit from glycine or a C3-unit derived from phosphoenolpyruvate as the remaining carbon(s) source (Miyakoshi et al., 1992; Berry and Abbott, 1978; Isono and Suhadolnik, 1976; and Ohnuki et al., 2003). These results have led to a unifying proposal that C-C bond formation occurs through an intermolecular aldol or aldol-type addition reaction following the generation of a nucleoside-5′-aldehyde that serves as the electrophile. As a consequence of this hypothesis, the biosynthesis of these compounds would require an oxidation step that would divert the canonical nucleoside into these specialized pathways.

The biosynthetic gene clusters for several of these compounds, including A-500359s from Streptomyces griseus SANK 60196 (Nonaka et al., 2009), A-503083s from Streptomyces sp. SANK 62799 (Funabashi et al., 2010), polyoxins from Streptomyces cacaoi (Chen, et al., 2009), nikkomycins from Streptomyces tendae Tü901 (Bruntner, et al., 1999), A-90289s from Streptomyces sp. SANK 60405 (Funabashi et al., 2010), caprazamycins from Streptomyces sp. MK730-62F (Kaysser et al., 2009), liposidomycins from Streptomyces sp. SN-1061M (Kaysser et al., 2010), muraymycins from Streptomyces sp. NRRL 30471 (Cheng et al., 2011), and tunicamycins from Streptomyces charteusis (Wyszynski et al., 2010 and Chen et al., 2010) have been cloned and sequenced, all within the past four years, excluding only the nikkomycin gene cluster, which was first identified in the 1990s. With the sole exception of the proposed gene cluster for tunicamycins, the gene clusters contain minimally one open reading frame encoding a protein with sequence similarity to enzymes of the Fe(II)-and α-ketoglutarate (αKG)-dependent dioxygenase superfamily (Schofield and Zhang, 1999; Hausinger, 2004; Bollinger Jr. et al., 2005). This enzyme superfamily, which includes taurine hydroxylase (TauD) and clavaminic acid synthase (CAS), are non-haem dioxygenases that couple the oxidative decarboxylation of typically αKG with oxidation—usually in the form of hydroxylation—of a second substrate. The latter, which is also called the prime substrate, includes a large and structurally diverse range of molecules (Hausinger, 2004).

LipL—a dioxygenase from this superfamily encoded within the A-90289 gene cluster—has been functionally assigned as an Fe(II)-dependent uridine-5′-monophosphate (UMP):αKG dioxygenase, thus establishing an enzymatic imperative for the formation of nucleoside-5′-aldehydes (Yang et al., 2011). LipL catalyzes a net dephosphorylation and oxidation of UMP to generate uridine-5′-aldehyde, which is the first intermediate in the biosynthesis of the unusual aminoribosyl moiety found in several C7-furanosyl nucleosides (Chi et al., 2011), and likely also serves to initiate assembly of the high-carbon sugar nucleoside scaffold (Figure 2). Herein is described the functional assignment and assay protocols for characterization of these new members of the dioxygenase superfamily.

Figure 2. Pathway involving LipL and homologous enzymes.

Figure 2

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Enzymes annotated in bold have been functionally characterized. The hypothetical intermediate from the LipL-catalyzed reaction is based on the hydroxylation mechanism established for TauD.

2. Methods

2.1. Bioinformatic analysis of LipL and homologous dioxygenases

Iron binding by the non-haem, αKG dioxygenase superfamily is mediated by the side-chains of Asp/Glu and His residues. The conserved and essential motif is HX1D/EXnH, wherein Xn can vary from 40 to 153 amino acids, although the spacing between ligand 2 (D/E) and ligand 3 (H) tends to be either approximately 50 or 140 residues, as previously noted by Hausinger (2004).

  1. Align the proteins using a ClustalW program with LipL as the reference protein (accession no. BAJ05888). An example of a sequence alignment of LipL with CapA involved in the biosynthesis of A-503083s is shown in Figure 3.

  2. Confirm the presence of the Fe(II)-binding motif that is essential for catalysis. In addition, identify the number of residues that constitute the spacer region between the Fe(II)-binding Ligands 2 and 3. Similarly to TauD (153 residues) and CAS (132 residues), the UMP:αKG dioxygenases have relatively long spacers: LipL, Cpz15 in the biosynthesis of caprazamycins, and LpmM in the biosynthesis of liposidomycins with 140 residues; Mur16 in the biosynthesis of muraymycins with 138 residues; and CapA and ORF7 in the biosynthesis of A-500359s with 139 residues.

Figure 3. Sequence analysis of LipL.

Figure 3

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The sequence for LipL is aligned with CapA, the homologous enzyme involved in the biosynthesis of A-503083s. Conserved regions are boxed and contain the residues critical for Fe(II) binding (▼).

2.2. Cloning and heterologous expression

The following protocol has been successfully used to express a number of genes from various actinomycetes. This cloning strategy also renders subsequent subcloning experiments relatively easy and straightforward (if necessary), since the entire gene along with the engineered codons encoding the His6-tag can be excised from the plasmid by NdeI and a second restriction enzyme, usually HindIII, BamHI, or EcoRI, for ligation into pUWL201 or a related Streptomyces expression vector.

  1. Design primers for PCR-amplification of the gene for insertion into the pET-30Xa ligation-independent cloning vector provided by Novagen (Madison, WI).

  2. Perform a PCR using the Expand Long Template PCR System from Roche (Indianapolis, IN) with Buffer 2 provided by the manufacturer. A typical reaction consists of 10 ng of template DNA, 200 μM each primer, 250 μM dNTPs, 5% DMSO, and 2.5 U of the DNA polymerase per 50 μL reaction. The thermocycler program includes an initial hold at 95 °C for 2 min followed by 30 cycles of 95 °C for 10 s, 56 °C for 15 s, and 68 °C for 75 sec.

  3. Purify the DNA fragment of the expected size by 1% agarose gel electrophoresis with ethidium bromide staining, recovering the PCR product using a commercial kit such as the Wizard SV Gel and PCR Clean-Up System from Promega (Madison, WI).

  4. Prepare the DNA overhangs using T4 DNA polymerase, following the protocol provided by Novagen, anneal with the pET-30Xa vector, and introduce by transformation into NovaBlue cells. Following identification of positive clones by colony PCR or restriction digests of the purified plasmid, the PCR-amplified DNA is sequenced to confirm its identity.

  5. Introduce the plasmid into E. coli BL21(DE3) by transformation and grow the recombinant strain in LB medium supplemented with 30 μg/mL kanamycin. Following inoculation of 500 mL of LB containing 30 μg/mL kanamycin in a 2.5-L Erlenmeyer flask, grow the recombinant strain at 18 °C with 250 rpm for ~ 9 hr. When the OD600 is ~ 0.5, add IPTG to a final concentration of 0.1 mM.

  6. Harvest the cells 14–16 hours after IPTG-induction and flash-freeze using liquid nitrogen for storage at −80 °C until needed.

  7. Thoroughly resuspend cells to a density of 200 mg/mL in 100 mM Tris-HCl pH 8.0 and 300 mM KCl, and store the suspension on ice for 10 min.

  8. Lyse the cells by one pass through a French press at 15,000 psi, and immediately centrifuge at 18,000 rpm for 30 minutes to remove the cell debris.

  9. Filter the supernatant with a low-protein binding, 0.45-μm syringe filter such as HPF Millex®-HV from Millipore (Billerica, MA), and load onto a column containing Ni-NTA resin from Qiagen (Valencia, CA) using approximately 1 mL resin per 1 g of dry cell mass. The Ni-NTA is pre-equilibrated with 10 column volumes of lysis buffer.

  10. Wash the column with 10 column volumes of lysis buffer followed by 10 column volumes of lysis buffer containing 20 mM imidazole.

  11. Elute the protein with 6 column volumes of 100 mM HEPES pH 7.5, 300 mM KCl, and 200 mM imidazole. Following ultrafiltration to reduce the volume to less than 2.5 mL, the purified protein is desalted into 100 mM HEPES pH 7.5, 50 mM KCl using PD-10 columns from GE Healthcare (Piscataway, NJ), and the resulting solution re-concentrated to ~ 500 μL prior to adding glycerol (final 40%) for storage at −20 °C.

  12. Assess solubility and purity by SDS-PAGE using 12% acrylamide. To serve as a control for ascertaining activity of the dioxygenase superfamily, the E. coli tauD can be cloned and expressed in an identical manner to yield soluble, active protein as shown in Figure 4 (Yang et al., 2011).

Figure 4. Purification of recombinant dioxygenases from E. coli.

Figure 4

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(A) SDS-PAGE of purified His6-LipL (expected 38.2 kD). (B) SDS-PAGE of purified His6-EcTauD (expected 37.4 kD). The engineered N-terminus His-tag contributes approximately 5 kD to the native molecular mass. This figure was originally published by Yang et al. (2011).

2.3. SEC-RI-MALLS-ICP-MS to determine metalloprotein stoichiometry

Fe(II) is weakly bound by the HX1D/EXnH motif and, depending on the isolation conditions and inherent properties of the protein, the recombinant enzyme can co-purify with varying levels of Fe(II). Inductively coupled plasma mass spectrometry (ICP-MS) is a powerful online chromatographic detector for identifying and quantifying metals such as iron and heteroatoms in proteins (Unrine et al., 2006; Unrine et al., 2007), which can be coupled on the front-end with size-exclusion chromatography (SEC) to remove unbound metal and protein aggregates under relatively mild conditions at physiological ionic strength and pH (Yang et al., 2011). In order to calibrate the ICP-MS, purified metalloproteins of known metal stoichiometry are used; hemoglobin is a good standard for the detection of iron. Metal-free chromatography components are utilized to minimize background concentrations or binding of exogenous metals to the proteins. For example, stainless steel tubing is replaced with polyethyl ether ketone (PEEK) tubing, and metal-free SEC columns are used. The ICP-MS technique is additionally coupled to refractive index (RI) and multi-angle laser light scattering detection (MALLS) for determining the protein concentration and molar mass, and hence allowing for the calculation of iron bound per unit of protein (Wyatt, 1993).

  1. Dissolve or dilute the unknown protein and protein standards to ~ 1 mg/mL in a mobile phase such as 150 mM NH4NO3 pH 7.2. In contrast to phosphate buffers, NH4NO3 is compatible with ICP-MS and is less likely to form problematic, insoluble metal complexes. Prior to injection, the protein sample is filtered using a 0.2-μm regenerated cellulose syringe filter to remove large aggregates and particulates.

  2. Equilibrate a high-resolution SEC column that has a selectivity within the range of expected molar mass of the protein, usually 600 kDa-5 kDa, with 3 column-volumes of mobile phase. An inline 0.1-μm hydrophilic fluoropolymer filter and a degasser between the pump and the column is used to avoid particles or gas bubbles which interfere with the light-scattering signal.

  3. Ensure that the MALLS detector is properly calibrated and normalized and the RI detector is properly calibrated. Additionally, the ICP-MS is optimized for the detection of iron or other metal of interest. The outlet of the column is directed first to the MALLS, followed sequentially by the RI detector and ICP-MS, using a splitter to match the optimal flow rate of the column with the optimal flow rate of the ICP-MS.

  4. Elute the standards and sample, choosing an appropriate range of protein concentration for the standards.

  5. Integrate the RI signal for the peaks representing the aggregated protein and calculate the mass of the protein assuming a dN/dC value of 0.185 if the actual dN/dC value is not known (Wyatt, 1993). The MALLS peak is integrated to determine the molar mass using a partial Zimm plot or an appropriate model for the molecule of interest (Wyatt, 1003; Zimm, 1945). Finally, signal intensity for the m/z of interest on the ICP-MS is integrated to calculate metal content.

  6. Calculate the number of moles of protein based on the total mass of protein and molar mass in each peak for the standards with known metal stoichiometry. A calibration curve of peak area versus moles of metal atoms is constructed from the ICP-MS data using least squares regression analysis.

  7. Determine the molar mass and mass of the unknown sample using the MALLS and RI data. This calculated molar value is compared to the number of moles of metal atom of interest from the corresponding peak in the ICP-MS chromatogram using the calibration curve. Using this methodology, LipL was isolated with 14 ± 2% mol iron per mol of protein, thus explaining why some activity is observed without exogenously added FeCl2 (Yang, 2011).

2.4. Activity Assays

The Fe(II)-dependent αKG:UMP dioxygenases utilize three substrates—UMP (or prime substrate), αKG, and O2—to generate four products, uridine-5′-aldehyde, phosphate, succinate, and CO2. Any of these reactants/products can be monitored to detect enzyme activity, and we routinely use the detection of three products as shown in Figure 5. For routine tests of activity, the reaction mixtures contain 50 mM HEPES pH 7.5, 1 mM UMP, 1.5 mM αKG, 200 μM ascorbic acid, 100 μM FeCl2, and 100 nM LipL unless noted herein. We typically use 50 mM HEPES pH 7.5 as a buffer in activity assays for HPLC since amine-containing buffers such as TRIS form an imine with the aldehyde product. Additionally, phosphate buffers and salts are avoided since phosphate is a product and inhibitor of the forward reaction.

Figure 5. Reaction and assay development for LipL.

Figure 5

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Three of the four products generated by the LipL-catalyzed reaction can be monitored using end-point (Malachite Green and HPLC) or continuous measurements (UV/Vis spectroscopy with an enzyme-coupled reaction).

2.4.1. Detection of succinate using an enzyme-coupled reaction

Most enzymes of the non-haem, Fe(II)-dependent dioxygenase superfamily incorporate one O atom from molecular oxygen into αKG to form CO2 and succinate. Thus an enzyme-coupled reaction leading to the oxidation of NADH has been developed that takes advantage of succinate formation (Luo et al., 2006). Several dioxygenases that generate succinate via oxidative decarboxylation of αKG often do so at reduced rates in the absence of the prime substrate.

  1. Prepare the appropriate solutions for performing the succinic acid assay procedure according to the protocol provided by Megazyme International (Wicklow, Ireland) except for the following changes: (i) 0.5 M HEPES or TRIS buffer replacing solution 1; (ii) only 1/10 volume is used per reaction; and (iii) αKG, FeCl2, ascorbic acid, and UMP (optional) are included, and the amount of water added is adjusted accordingly.

  2. Initiate the reaction by adding 100–500 nM of dioxygenase, and monitor the activity by following the loss of absorbance at 340 nm over 10 min at 30 °C.

  3. Calculate the specific activity using ε340 nm = 6,220 M−1 for β-NADH and compare the activity with and without UMP to obtain a relative specific activity. The negative control should contain all of the components except αKG.

2.4.2. HPLC analysis

Both the substrate and product are very hydrophilic, which complicates analysis of the reaction using typical C18 reverse-phase chromatography conditions. However, ion-pairing chromatography offers an adequate alternative for separating these compounds. Unfortunately, the product aldehyde is too unstable for column purification under a variety of conditions, so co-elution with authentic standard and LC-MS, the latter using standard reverse-phase chromatography conditions, are essential for confirming its identity.

  1. Synthesize the product uridine-5′-aldehyde using a routine, 3-step procedure as previously described (Yang et al., 2011).

  2. Prepare a 0.5-mL reaction containing the standard components and initiate by adding LipL, incubating at 30 °C. Negative controls consist of the reaction mixture either without enzyme or without αKG.

  3. At different time-points following the addition of the enzyme, pipet an 80 μL aliquot into an ultrafiltration device (3,000-molecular weight cutoff).

  4. Centrifuge in a bench-top microcentrifuge at maximum speed for 3 minutes.

  5. Recover the flow-through and analyze the sample using an HPLC equipped with a C18 reverse phase column (250 × 4.6 mm, 5 mm) using a mobile phase of 40 mM phosphoric acid-triethylamine (pH 6.5) with a linear gradient to 90% acetonitrile in 40 mM phosphoric acid-triethylamine (pH 6.5). The flow-rate is kept constant at 1.0 mL/min and nucleosides are detected at 260 nm using a photodiode array detector (Figure 6).

Figure 6. HPLC analysis to detect UMP:αKG dioxygenase activity.

Figure 6

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Time-course analysis of the LipL reaction using HPLC with a diode array detector for separation and detection. (○), UMP; (●), uridine-5′-aldehyde; A260, absorbance at 260 nm.

2.4.3 Malachite-green binding assay

The detection of phosphate as an enzyme product is well established for assaying the activity of phosphatases (Fisher and Higgens, 1994), and as such several vendors sell malachite-green (MG) phosphate detection kits. We have successfully employed the colorimetric-based SensoLyte™ MG Phosphatase Assay Kit from AnaSpec, Inc. (Fremont, CA).

  1. Incubate a 100 uL solution of 50 mM HEPES pH 7.5, 1 mM UMP, 1.5 mM αKG, 200 μM ascorbic acid and 100 μM FeCl2 for 1 min at 30 °C and initiate the reaction by adding LipL (100 nM final) using the pipettor for mixing.

  2. Remove 80 μL of the reaction mixture and add to 20 uL of the MG reagent that was previously dispensed into individual wells in a 96-well format. Mix well and incubate at room temperature for 10 minutes. The MG reagent contains 1 M sulfuric acid, which was shown to terminate the reaction by analyzing the formation of uridine-5′-aldehyde at different time-points following the addition of MG reagent, using HPLC as the method of detection.

  3. Measure the absorbance at 620 nm using a microplate reader. A standard curve is developed using phosphate stocks of 0.8, 1.6, 3.2, 6.7, 12.5, 25, and 50 μM. When varying the concentration of FeCl2 or ascorbic acid, prepare blanks at the respective concentrations that do not include the enzyme.

  4. For single-substrate kinetic analysis, conduct reactions for 3 min (< 10% product formation) and vary αKG (5 μM–3 mM) with saturating UMP (2 mM) or vary UMP (5 μM–1 mM) with saturating αKG (1 mM), again using a sample without enzyme as a blank/control. The simplicity of the assay allows for several replicates to be performed, although it is important to maintain approximately the same developing time following the addition of the MG reagent.

Summary

The functional characterization of LipL has established a dioxygenase-mediated strategy as one potential mechanism for the formation of a nucleoside-5′-aldehyde from the monophosphorylated nucleotide. Other mechanisms do exist; for example, biosynthesis of pacidamycins, which contain a 5′-amino-5′-deoxyuridine core, occurs through the formation of uridine-5′-aldehyde from uridine catalyzed by a flavin-dependent oxidoreductase, Pac11 (Ragab et al., 2011). Additionally, the nikkomycin and polyoxin gene clusters contain multiple gene products resembling other enzymes of the Fe(II)- and αKG-dependent dioxygenase superfamily. However, these enzymes (such as NikI and NikM in nikkomycin biosynthesis) have closer sequence similarity to prolyl-4-hydroxylases, which contain a short spacer (HKDX55H for NikI and HTDX56H for NikM), suggesting a unique activity. Furthermore, UMP is first converted to 3′-enoylpyruvyl-UMP during nikkomycin biosynthesis (Ginj et al., 2005), so despite the strong probability that the biosynthesis proceeds through a 5′-aldehyde, the role of these dioxygenases in forming the high-carbon furanoside remains to be established. Nonetheless, Fe(II)- and αKG-dependent enzymatic hydroxylation vicinal to a phosphate group either as a strategy to form nucleoside-5′-aldehyde such as the function of LipL or to release phosphate—the latter of which has been speculated as a potential mechanism for phosphate salvage (Hausinger, 2004)—can be readily identified and characterized, in part utilizing the described protocols.

Acknowledgments

This work is supported by grants to S.G.V.L. from the American Cancer Society (IRG-85-001-19), Kentucky Science and Technology Corporation, and National Institutes of Health (AI087849).

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