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. Author manuscript; available in PMC: 2019 Mar 16.
Published in final edited form as: Methods Enzymol. 2018 Mar 16;605:253–265. doi: 10.1016/bs.mie.2018.01.031

Preparation and Characterization of Tetrabromopyrrole Debrominase From Marine Proteobacteria

Jonathan R Chekan *, Bradley S Moore *,†,1
PMCID: PMC6211843  NIHMSID: NIHMS993374  PMID: 29909826

Abstract

While halogenases have been studied for decades, the first natural product dehalogenase was only recently described. This bacterial enzyme, Bmp8, catalyzes the reductive debromination of 2,3,4,5-tetrabromopyrrole to form 2,3,4-tribromopyrrole as part of the biosynthesis of pentabromopseudilin, a marine natural product. Bmp8 is hypothesized to utilize a catalytic mechanism analogous to the important human thyroid hormone deiodinase enzyme family, potentially enabling Bmp8 to serve as model system to study this conserved mechanism. Herein, we describe a method for the soluble expression and purification of Bmp8. Furthermore, we detail activity assay protocols to quantify both consumption of the tetrabromopyrrole substrate and formation of the tribromopyrrole product. These methods will enable further study of this unusual enzyme and its catalytic mechanism.

1. INTRODUCTION

Halogenation biochemistry is an abundant and important transformation in natural product biosynthesis (Agarwal et al., 2017). Present in all domains of life, and particularly prevalent in marine environments, halogenation reactions have been observed to utilize all the commonly occurring halide ions (Gribble, 2010). The halogen atoms present on a final natural product chemical are often critical for its biological activity, and organohalogens are viewed as attractive leads for drug development in the pharmaceutical industry (Hardegger et al., 2011; Lu et al., 2012; Xu et al., 2011). While the enzymatic installation of a halogen atom has been well studied in many natural product systems (Agarwal et al., 2017), removal of a halogen is much less understood. Dehalogenation has generally been examined in the context of bioremediation of polyhalogenated humanmade (anthropogenic) compounds that accumulate in the environment as persistent toxins (Jugder, Ertan, Lee, Manefield, & Marquis, 2015). These dehalogenases utilize diverse mechanisms and it is unknown whether they evolved to degrade human-produced chemicals or naturally occurring halogenated compounds (Vetter, 2006).

In spite of the extensive research on halogenation, only a single recently characterized bacterial natural product dehalogenase, Bmp8, has been identified to naturally function in a biosynthetic pathway (El Gamal, Agarwal, Rahman, & Moore, 2016). This enzyme was uncovered upon the elucidation of the biosynthetic pathway of pentabromopseudilin, a polybrominated marine antibiotic from marine gammaproteobacteria Pseudoalteromonas luteoviolacea and Marinomonas mediterranea (Fig. 1) (Agarwal et al., 2014; El Gamal, Agarwal, Diethelm, et al., 2016). Early characterization of the pathway revealed the presence of a 2,3,4,5-tetrabromopyrrole intermediate (El Gamal, Agarwal, Diethelm, et al., 2016). Notably, this bacterial compound has coral settlement activity that may assist in the growth and maintenance of coral reefs (Sneed, Sharp, Ritchie, & Paul, 2014). Tetrabromopyrrole appeared to be an over brominated compound because only three bromines are present on the final pyrrole moiety (Fig. 1), and the final enzyme in the pathway utilized the tribromopyrrole species as a substrate (Agarwal et al., 2014). A more detailed exploration of the pentabromopseudilin biosynthetic gene cluster enabled expression and purification of Bmp8, a 21kDa protein with limited sequence similarity to the enzyme AhpD (El Gamal, Agarwal, Rahman, et al., 2016). While the physiological function of AhpD is still unclear, this Mycobacterium tuberculosis protein has been shown to possess both disulfide reductase and alkylhydroperoxidase activities (Bryk, Lima, Erdjument-Bromage, Tempst, & Nathan, 2002; Hillas, Soto Del Alba, Oyarzabal, Wilks, & Ortiz De Montellano, 2000; Nunn, Djordjevic, Hillas, Nishida, & Ortiz De Montellano, 2002). This similarity seemed to suggest that Bmp8 could catalyze reductive debromination chemistry via a thioredoxin-like reaction mechanism. Subsequent activity assays demonstrated that Bmp8 could catalyze the conversion of tetrabromopyrrole to tribromopyrrole, the substrate needed for formation of the final pentabromopseudilin natural product (Figs. 1 and 2) (El Gamal, Agarwal, Rahman, et al., 2016). Further exploration of Bmp8 revealed that it was cofactor-independent and utilized a conserved cysteine pair to catalyze the debromination (Fig. 2) (El Gamal, Agarwal, Rahman, et al., 2016). While the details of the mechanism are unclear, Bmp8 ends the reaction with the formation of a disulfide bond that needs to be reduced before another turnover can be completed (Fig. 2) (El Gamal, Agarwal, Rahman, et al., 2016). Chemical reductants can be used in vitro, but the physiological external reductant is currently unknown.

Fig. 1.

Fig. 1

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Biosynthesis of the antibiotic pentabromopseudilin.

Fig. 2.

Fig. 2

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Catalytic cycle of Bmp8.

Not only is Bmp8 interesting from a microbial biosynthetic viewpoint, but Bmp8 may be able to offer insight into human iodothyronine deiodinases. These human enzymes are responsible for maintaining the proper concentrations of the different thyroid hormones in the human body (Gereben et al., 2008). By controlling the number and location of iodines found on the thyroid hormones, the deiodinases can influence the metabolism levels of the entire body (Gereben et al., 2008). Therefore, misfunction of these deiodinases can lead to hyperthyroidism or hypothyroidism conditions (Gereben et al., 2008; Maia, Goemann, Meyer, & Wajner, 2011). One family of deiodinases, the selenocysteine thyroid hormone deiodinases (Dios), employ a catalytic pair of selenocysteine and cysteine residues for the deiodination activity (Agarwal et al., 2017; Schweizer, Schlicker, Braun, Kohrle, & Steegborn, 2014; Schweizer & Steegborn, 2015), very analogous to Bmp8’s use of two cysteines. There has been much debate on the nature of the catalytic mechanism of Dios (Schweizer & Steegborn, 2015), which could be important for the development of drugs that target Dios and modulate their activity. Unfortunately, studying Dios in an in vitro setting is often challenging both because they contain transmembrane helices, and they need the rare amino acid selenocysteine to be incorporated. Therefore, the ease of Bmp8 expression and manipulation may enable its use to probe the nature of this enzymatic dehalogenation mechanism.

In this chapter, we focus first on the expression and purification of Bmp8. We next describe an optimization of the Bmp8 activity assay protocol with special attention paid to designing reaction conditions that maximize the stability of tetrabromopyrrole.

2. HETEROLOGOUS EXPRESSION AND PURIFICATION OF Bmp8

Early attempts to heterologously express Bmp8 in Escherichia coli cells resulted in either no expression or insoluble proteins. During the course of probing the entire pentabromopseudilin biosynthetic pathway, the Bmp8 and the Bmp1 thioesterase (TE) domains were serendipitously expressed together. In the pathway, Bmp1 works in conjunction with Bmp2 to produce tetrabromopyrrole by brominating and offloading the acyl carrier protein-tethered tribromopyrrole (Fig. 1) (El Gamal, Agarwal, Diethelm, et al., 2016). Unexpectedly, this Bmp8 and Bmp1(TE) coexpression methodology produced soluble Bmp8. These two proteins did not form a stable complex because Bmp8 is retained on the initial His6 affinity column whereas Bmp1(TE) is mostly present in the wash step (El Gamal, Agarwal, Rahman, et al., 2016). Therefore, it appears that Bmp1(TE) may act as a chaperone or scaffold to assist with proper folding of Bmp8. Once correctly folded, Bmp8 remains stable and soluble.

2.1. Gene Cloning and Expression Strain Preparation

Both the bmp8 and bmp1(TE) genes were obtained by PCR from M. mediterranea MMB-1 DNA. The bmp8 gene was cloned into a pET28a vector using the NdeI/XhoI cut sites. This created an expression construct with a N-terminal His6 tag and kanamycin resistance. The bmp1(TE) gene was cloned into the multiple cloning site 2 (MCS2) of a pCDFDuet vector. This produced a spectinomycin resistance construct that lacked any additional tag upon bmp1(TE) expression. These two plasmids were cotransformed into E. coli BL21(GOLD) bacteria producing cells with both kanamycin and spectinomycin resistance. In all the experiments, streptomycin can be used in place of spectinomycin.

2.2. Bmp8 Overexpression

  1. From a glycerol stock of the cotransformed E. coli BL21(GOLD) cells, inoculate 10mL of Lysogeny Broth (LB) media supplemented with 50μg/mL of both kanamycin and spectinomycin. Incubate overnight at 37°C and shake at approximately 250rpm.

  2. Inoculate 1L of sterile Terrific Broth (TB) in a 2.8L Erlenmeyer flask with 2mL of the 10mL overnight culture. The media should again be supplemented with 50μg/mL of both kanamycin and spectinomycin.

  3. Incubate the 1L culture at 37°C and shake at 200rpm until OD600 reaches 0.6 (~4.5h).

  4. Adjust the temperature of the shaking incubator to 18°C and continue shaking the flask for one additional hour.

  5. Induce protein expression with a final isopropyl-β-D-thiogalactopyranoside (IPTG) concentration of 0.3mM.

  6. Continue incubation at 18°C and shaking at 200rpm overnight (~18h).

  7. Pellet the cells at 7000 × g for 7min and dispose of the spent media.

  8. Resuspended the entire cell pellet in 25mL of cold buffer containing 20mM Tris pH 8.0, 500mM NaCl, and 10% glycerol.

  9. Cells can be stored at ‒80°C or immediately lysed for Bmp8 purification.

2.3. Bmp8 Purification

  1. Thaw the frozen cell pellet and sonicate on ice to lyse cells. A Qsonica 6mm tip at 40% amplitude for 12 cycles of 15s on and 45s off was sufficient to lyse the cells.

  2. Centrifuge the cell lysate at 15,000 × g for 25min at 4°C to pellet insoluble cell debris.

  3. Equilibrate a 5mL His-Trap FF column (GE Healthcare Life Sciences) with 30mL of buffer A (20mM Tris pH 8.0, 1M NaCl, and 30mM imidazole).

  4. Load the supernatant onto the 5mL His-Trap FF column at a rate a 2mL/min using an ÄKTA FPLC system (GE Healthcare Life Sciences).

  5. Once loaded, wash the column with 40mL of buffer A to remove weakly bound proteins.

  6. Elute Bmp8 with a 40mL linear gradient starting with 100% buffer A and ending with 100% buffer B (20mM Tris pH 8.0, 1M NaCl, and 250mM imidazole). Use a flow rate of 2mL/min and collect 5mL fractions.

  7. Run a sample of the initial supernatant, pellet, flow through, wash, and each of the fractions on a 12% SDS-PAGE gel to assess purity of the Bmp8 fractions (Fig. 3A).

  8. Collect and combine the most pure fractions and save a small aliquot (~20μL) as an uncut standard.

  9. To remove the His6 affinity tag, add 120units of thrombin to the combined fractions and dialyze overnight (~16h) against 1L of cold buffer containing 50mM KCl, 20mM Tris pH 8.9, and 2mM DTT using a 14kDa molecular weight cut-off dialysis bag.

  10. Run the cut and uncut samples on a 12% acrylamide SDS-PAGE gel to confirm complete removal of the tag by thrombin (Fig. 3B).

  11. Make 250mL of cold buffer C (20mM Tris pH 8.9 and 3mM DTT) and buffer D (20mM Tris pH 8.9, 1M KCl, and 3mM DTT).

  12. Load the Bmp8 sample onto a 5mL strong anion exchange Q FF column (GE Healthcare Life Sciences) that has been preequilibrated with 95% buffer C and 5% buffer D. Complete all anion exchange purification steps using a flow rate of 2mL/min.

  13. Wash the column with 10mL of 5% buffer D to remove any weakly bound protein.

  14. While collecting 3mL fractions, elute Bmp8 using an 80mL linear gradient starting with 5% buffer D and ending with 70% buffer D. Bmp8 should begin to elute at approximately 25% buffer D. Run a 12% acrylamide SDS-PAGE gel to determine the purity of the fractions (Fig. 3B). Combine the pure fractions and concentrate with a Millipore Amicon Ultra-4 10kDa filter if desired.

  15. Load 2mL of Bmp8 onto a HiLoad 16/60 Superdex 75 prep grade column (GE Healthcare Life Sciences) preequilibrated with Bmp8 storage buffer (20mM Tris pH 8.0, 300mM KCl, and 10% glycerol). Collect 3mL fractions at a flow rate of 1mL/min. Bmp8 elutes at approximately 67mL, most consistent with a dimeric oligomerization state. This size exclusion column step further purifies Bmp8 while also removing DTT that would interfere with subsequent activity assays.

  16. Combine the Bmp8 elution peak and concentrate in a Millipore Amicon Ultra-4 10kDa molecular weight cut-off filter to approximately 500μL (~13mg/mL). 50μL aliquots can be frozen in liquid nitrogen or dry ice then stored at ––80°C for later use (Fig. 3C).

Fig. 3.

Fig. 3

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SDS-PAGE gels of (A) His6 affinity column fractions, (B) anion exchange column fractions, and (C) collected and concentrated gel filtration peak. The gels were loaded as follows: (A) 1-supernatant, 2-pellet, 3-flow through, 4-wash, M-marker, 5–14-elution fractions; (B) 1-thrombin cut, 2-uncut, 3-cut, 4-uncut, M-marker, 5–8-elution peak; and (C) M-marker, 1-concentrated Bmp8 after collecting gel filtration column peak. The 20 and 30kDa bands of the Fisher BioReagents EZ-Run Rec Protein Ladder are indicated in each gel. The Bmp1(TE) band can be observed just above the 30kDa band. Expected protein sizes: His6-Bmp8: 23.4kDa, Cut Bmp8: 21.6kDa, and Bmp1(TE): 35.7kDa.

3. Bmp8 ACTIVITY ASSAY

3.1. Assay Methodology

  1. All activity assays were performed at a 400μL scale in a 1.5mL microcentrifuge tube.

  2. Set up Bmp8 assays in a reaction mixture composed of 50mM KCl, 50mM Tris pH 8.0, 500μM glutathione, and 20μM Bmp8. Initiate reactions with 100μM tetrabromopyrrole and incubate at 30°C for 30min. The optimization of these conditions is discussed in Section 3.4.

  3. Extract both the tetrabromopyrrole substrate and tribromopyrrole product away from the reducing agent and Bmp8 by adding 800μL ethyl acetate to the reaction.

  4. Vortex the mixture to ensure thorough extraction and centrifuge for 5min at 15,000rpm to separate the water and ethyl acetate phases.

  5. Transfer 700μL of the ethyl acetate (top layer) into a fresh 2mL microcentrifuge tube.

  6. Remove the ethyl acetate with a centrifugal evaporative concentrator (SpeedVac/miVAC type system). Using a miVac DNA system (SP Scientific) at 30°C removes all the ethyl acetate in approximately 35min.

  7. Resuspend the residue in 80μL of 50% methanol. Higher concentrations of organic solvents will lead to peak broadening during HPLC analysis.

3.2. LC/MS-Based Reaction Evaluation

  1. Set up the following method to observe Bmp8 catalyzed conversion of tetrabromopyrrole to tribromopyrrole: 20min at 52% solvent B, gradient of 52% to 95% solvent B over 1min, 95% solvent B for 3min, gradient of 95% solvent B to 52% solvent B over 1min, and 2min at 52% solvent B. Solvent A is HPLC grade water +0.1% formic acid and solvent B is HPLC grade acetonitrile +0.1% formic acid. The tribromopyrrole and tetrabromopyrrole have expected M – 1 masses of 301.8 and 381.7m/z, respectively, in the negative ion mode.

  2. Inject 5 μL of the reaction onto a Phenomenex Luna 5μm C18(2) 100Å 150× 4.6mm2 column using the described method (Fig. 4).

Fig. 4.

Fig. 4

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LC/MS analysis of the Bmp8 reaction. The tetrabromopyrrole and tribromopyrrole extracted base peak chromatograms from both the Bmp8 reaction (bottom traces) and no enzyme control (top traces) were extracted using 381.7m/z (orange traces) and 301.8m/z (blue traces), respectively. This data were produced using an Agilent 1260 infinity HPLC and Bruker amaZon ion trap mass spectrometer.

3.3. HPLC Substrate and Product Quantification

  1. Set up the following method to analyze the amount of tetrabromopyrrole remaining and tribromopyrrole formed: 20min at 52% solvent B, gradient of 52%–95% solvent B over 1min, 95% solvent B for 3min, gradient of 95% solvent B to 52% solvent B over 1min, and 2min at 52% solvent B. Solvent A is HPLC grade water +0.1% trifluoroacetic acid (TFA) and solvent B is HPLC grade acetonitrile +0.1% TFA. If the TFA is omitted, the elution peaks are broadened. Monitor the presence of both tribromopyrrole and tetrabromopyrrole at a UV absorbance of 220nm.

  2. Inject 50μL of the 80μL reaction onto a Phenomenex Luna 5μm C18(2) 100Å 150 × 4.6mm2 column using the described method.

  3. Integrate the 220nm absorbance peaks at approximately 11.6 and 18.2min to determine the amount of tribromopyrrole and tetrabromopyrrole, respectively (Fig. 5). A pure sample of tribromopyrrole or tetrabromopyrrole can be used to create a standard curve to perform absolute quantification.

Fig. 5.

Fig. 5

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HPLC analysis of the Bmp8 catalyzed debromination of tetrabromopyrrole. UV absorption was monitored at 220nm in both the no Bmp8 reaction (black trace) and the Bmp8 reaction (red trace). This data were collected on an Agilent 1200 series HPLC.

3.4. Tetrabromopyrrole Stability Evaluation

The inherent instability of tetrabromopyrrole required a thorough optimization of the assay conditions to limit the nonenzymatic reductive debromination.

3.4.1. Reductant Choice

The catalytic mechanism of Bmp8 requires an external reductant to regenerate the active form of the enzyme (Fig. 2). As the native reductant is unknown, several chemical reductants were evaluated to determine which has the lowest nonenzymatic reactivity with tetrabromopyrrole. Reactions were completed with 50mM KCl, 20mM Tris pH 8.0, 100μM tetrabromopyrrole, and 500μM of either dithiothreitol (DTT), glutathione (GSH), or tris(2-carboxyethyl)phosphine (TCEP). The assay methodology described in Section 3.1 was utilized, and the reaction was monitored by the HPLC method described in Section 3.3. The results indicated that tetrabromopyrrole was most stable in GSH or TCEP and that DTT catalyzed the most nonenzymatic debromination (Fig. 6A). Subsequent reactions were performed with GSH as its biological origin suggests that it might be a more physiological reductant.

Fig. 6.

Fig. 6

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Nonenzymatic debromination of tetrabromopyrrole by (A) different reductants and in (B) different buffers systems. Tetrabromopyrrole and tribromopyrrole were quantified based on integration of the 220nm absorbance peaks and normalized against the maximal tetrabromopyrrole integration. Tetrabromopyrrole amounts are shown in solid bars and tribromopyrrole quantifications are in striped bars. These assays were completed in triplicate.

3.4.2. Buffer Choice

The stability of tetrabromopyrrole was also evaluated in the presence of different buffering agents and pHs. HEPES and Tris (both Good’s buffers) were tested at pH 7.0, 7.5, and 8.0 for tetrabromopyrrole degradation. Reactions were performed with 50mM KCl, 500μM glutathione, 100μM tetrabromopyrrole, and 20mM of the desired buffer and pH. The assays were completed as described in Section 3.1 and monitored for substrate stability as described in Section 3.3. Tetrabromopyrrole appeared to decay in the HEPES buffer in a pH-dependent manner, with the least degradation occurring at pH 7.0 and the most at pH 8.0 (Fig. 6B). Conversely, the tetrabromopyrrole was more stable in the Tris buffers and pH did not appear to alter the stability significantly. These results align with previous observations that HEPES is able to form radicals and is not an ideal buffer for redoxsensitive assays (Kirsch, Lomonosova, Korth, Sustmann, & de Groot, 1998). Based on these results, 20mM Tris pH 8.0 was selected as buffer for downstream activity assays.

4. CONCLUSIONS

Bmp8 has been shown to catalyze the debromination of the coral settlement cue tetrabromopyrrole to the pentabromopseudilin precursor tribromopyrrole (El Gamal, Agarwal, Rahman, et al., 2016). Difficulties in heterologous expression revealed that the coexpression of Bmp1(TE) was required for soluble and stable production of Bmp8. The inherent instability of the tetrabromopyrrole substrate required extensive optimization to yield a quantitative assay that can monitor the reductive debromination activity of Bmp8. These procedures lay the groundwork for further mechanistic studies of this unusual natural product biosynthetic enzyme and its potential use as a model system for human iodothyronine deiodinases.

ACKNOWLEDGMENTS

We thank the NSF (OCE-1313747) and NIEHS (P01-ES021921) through the Oceans and Human Health program for financial support.

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