Five Fatty Aldehyde DehydrogenaseEnzymes from Marinobacter andAcinetobacter spp. and StructuralInsights into the Aldehyde BindingPocket

Jonathan H. Bertram,a Kalene M. Mulliner,a Ke Shi,d Mary H. Plunkett,b

Peter Nixon,a Nicholas A. Serratore,c Christopher J. Douglas,c Hideki Aihara,d

Brett M. Barneya,b

Department of Bioproducts and Biosystems Engineeringa and Biotechnology Institute,b University ofMinnesota, St. Paul, Minnesota, USA; Department of Chemistryc and Department of Biochemistry, MolecularBiology and Biophysics,d University of Minnesota, Minneapolis, Minnesota, USA

ABSTRACT Enzymes involved in lipid biosynthesis and metabolism play an impor-tant role in energy conversion and storage and in the function of structural compo-nents such as cell membranes. The fatty aldehyde dehydrogenase (FAldDH) plays acentral function in the metabolism of lipid intermediates, oxidizing fatty aldehydesto the corresponding fatty acid and competing with pathways that would further re-duce the fatty aldehydes to fatty alcohols or require the fatty aldehydes to producealkanes. In this report, the genes for four putative FAldDH enzymes from Marinobac-ter aquaeolei VT8 and an additional enzyme from Acinetobacter baylyi were heterolo-gously expressed in Escherichia coli and shown to display FAldDH activity. Five en-zymes (Maqu_0438, Maqu_3316, Maqu_3410, Maqu_3572, and the enzyme reportedunder RefSeq accession no. WP_004927398) were found to act on aldehydes rangingfrom acetaldehyde to hexadecanal and also acted on the unsaturated long-chainpalmitoleyl and oleyl aldehydes. A comparison of the specificities of these enzymeswith various aldehydes is presented. Crystallization trials yielded diffraction-qualitycrystals of one particular FAldDH (Maqu_3316) from M. aquaeolei VT8. Crystals wereindependently treated with both the NAD� cofactor and the aldehyde substratedecanal, revealing specific details of the likely substrate binding pocket for this classof enzymes. A likely model for how catalysis by the enzyme is accomplished is alsoprovided.

IMPORTANCE This study provides a comparison of multiple enzymes with the abil-ity to oxidize fatty aldehydes to fatty acids and provides a likely picture of how thefatty aldehyde and NAD� are bound to the enzyme to facilitate catalysis. Based onthe information obtained from this structural analysis and comparisons of specifici-ties for the five enzymes that were characterized, correlations to the potential rolesplayed by specific residues within the structure may be drawn.

KEYWORDS Marinobacter, Maqu_3316, decanal, wax ester, lipid biosynthesis

The fates of fatty compounds within the cell are a central aspect of lipid metabolism.Enzymes involved in the metabolism of fatty compounds can play important roles

in disease and have a biotechnological relevance for both the production of lipids orbiofuels and the degradation of oils that are released into the environment (1). Thebacterium Marinobacter aquaeolei VT8 was isolated from the head of an offshore oilwell near Vietnam, where it would be expected to be participating in the biodegrada-

Received 4 January 2017 Accepted 3 April2017

Accepted manuscript posted online 7 April2017

Citation Bertram JH, Mulliner KM, Shi K,Plunkett MH, Nixon P, Serratore NA, Douglas CJ,Aihara H, Barney BM. 2017. Five fatty aldehydedehydrogenase enzymes from Marinobacterand Acinetobacter spp. and structural insightsinto the aldehyde binding pocket. ApplEnviron Microbiol 83:e00018-17. https://doi.org/10.1128/AEM.00018-17.

Editor Rebecca E. Parales, University ofCalifornia—Davis

Copyright © 2017 American Society forMicrobiology. All Rights Reserved.

Address correspondence to Brett M. Barney,bbarney@umn.edu.

ENZYMOLOGY AND PROTEIN ENGINEERING

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tion of crude oil inadvertently released into the environment (2). In addition to oilbioremediation in the natural environment, this bacterium also produces a high-valuelipid, the wax ester, which has commercial significance and is used in a range ofcommodity products (3). M. aquaeolei VT8 is relatively easy to culture in the laboratory,and many of the enzymes obtained from this species show strong activity whenexpressed in heterologous hosts such as Escherichia coli, Saccharomyces cerevisiae, andSynechococcus elongatus (4–6). For this reason, M. aquaeolei VT8 has become a modelspecies for the study of lipid metabolism and an important source of enzymes forbiotechnological applications.

Our laboratory has an interest in studying the enzymes that take part in theproduction of high-value compounds associated with wax ester production, includingthose that reduce fatty acyl coenzyme A (CoA), fatty acyl-acyl carrier protein (ACP), orfatty aldehydes to fatty alcohols (7–10). Fatty alcohols are a specific substrate for thewax ester synthase (3, 11, 12). In addition to enzymes that reduce fatty aldehydes tofatty alcohols, there are also enzymes present in M. aquaeolei VT8 that could oxidize thefatty aldehyde back to a fatty acid, such as the fatty aldehyde dehydrogenase (FAldDH).A previous report characterized a FAldDH from another species that accumulates waxesters, Acinetobacter sp. strain M-1 (13). A BLAST search using the primary sequence ofthe FAldDH from Acinetobacter sp. strain M-1 revealed a large number of genes from M.aquaeolei VT8 that might also function as FAldDHs (13–15). These enzymes are ofparticular concern in biosynthetic strategies, because they could result in a futilepathway that would diminish the accumulation wax ester or other targeted products,especially for enzymes that utilize fatty aldehydes, including fatty aldehyde decarbony-lases or fatty acyl-CoA reductases and fatty aldehyde reductases (8–10, 16, 17). Oneparticular fatty aldehyde dehydrogenase (Maqu_3410) was highly transcribed duringwax ester accumulation in M. aquaeolei VT8 when grown on the simple carbohydratecitrate (CIT) or malate (8). For this reason, and because a thorough analysis of a varietyof putative FAldDHs from bacteria is lacking in the literature, we sought to isolate andcharacterize a range of these putative FAldDHs from M. aquaeolei VT8. Furthermore, weselected one enzyme for structural studies, and here we report the crystal structures ofthe Maqu_3316 FAldDH from M. aquaeolei VT8 in complex with the NAD� cofactor ora fatty aldehyde substrate.

RESULTSEnzyme activity and specificity with fatty aldehydes. The primary aim of this

research was to characterize the fatty aldehyde oxidation activities of several putativeFAldDH enzymes from Marinobacter aquaeolei VT8. An initial survey of the M. aquaeoleiVT8 genome (15) using the BLAST algorithm (14) and the previously reported FAldDHsequence from Acinetobacter sp. strain M-1 (13) revealed a large number of genes withsignificant amino acid sequence identity to this previously characterized FAldDH(GenBank accession no. AB042203). Five putative FAldDH genes from M. aquaeolei VT8and an additional gene from Acinetobacter baylyi were cloned and heterologouslyexpressed in Escherichia coli. Each gene was modified to incorporate a polyhistidine tag(at the N terminus) to allow rapid purification. The isolated proteins obtained followingmetal affinity chromatography purification and a subsequent desalting step are shownin Fig. 1. A band corresponding to the expected protein size was found for each of thesix proteins, although only two of the enzymes (Maqu_3316 and Maqu_3410) wereobtained as high-purity single bands based on this single-step purification. Three of theenzymes (Maqu_0607, Maqu_3572, and the enzyme reported under RefSeq accessionno. WP_004927398) showed minor contaminating or degradation bands, whileMaqu_0438 showed the greatest degree of degradation and the lowest purity. Somevariability was found for specific enzyme preparations, especially in relation to proteinstability and activity. For several enzymes, the activity in certain preparations was verylow, although the protein purity and quantities obtained were comparable from onepreparation to another. To a certain extent, some of this activity could be rescued bydegassing the protein preparation under an argon atmosphere and then adding a small

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quantity (5 �l) of �-mercaptoethanol to �8 ml of a solution containing the purifiedprotein and allowing it to stand for several minutes. This was most prominently foundfor Maqu_3410, which seemed to be particularly susceptible to this issue.

An initial assessment of the substrate specificities of the M. aquaeolei VT8 enzymesusing saturated, straight-chain aldehydes ranging from butanal (C4) to hexadecanal(C16) showed that four of the enzymes exhibited aldehyde dehydrogenase activity, andeach of these enzymes had unique substrate selectivity patterns (Fig. 2). We alsoperformed an alignment of these proteins with sequences of enzymes from the ProteinData Bank (PDB) that have been reported to have aldehyde dehydrogenase activity andgenerated a phylogenetic tree based on these alignments (Fig. 3). Maqu_0438 andMaqu_3572 clustered together in this tree based on their primary sequences and alsodisplayed similar substrate selectivity profiles (Fig. 2) that showed a broad, almostGaussian distribution for both proteins with straight-chain fatty aldehydes. Maqu_3316showed a more prominent specificity for decanal, with activity dropping considerablyfor both octanal and dodecanal versus decanal. Maqu_3316 also clustered separatelyfrom the other enzymes tested here (Fig. 3). Maqu_3410 showed a much moreconsistent level of activity across the entire range, with high selectivity for longer fattyaldehydes such as hexadecanal, and was the only enzyme from M. aquaeolei VT8 thatyielded a specificity profile that was similar to what was reported previously by Ishigeet al. for their FAldDH enzyme from Acinetobacter sp. strain M-1 (13). In addition to theM. aquaeolei VT8 FAldDHs, we also isolated a homologous FAldDH from A. baylyi(RefSeq accession no. WP_004927398), which had very high amino acid sequenceconservation with the sequence for the FAldDH enzyme from Acinetobacter sp. strainM-1. Maqu_3410 and the enzyme reported under RefSeq accession no. WP_004927398also clustered together based on their primary sequences (Fig. 3) and displayed similartrends for their substrate selectivity profiles (Fig. 2). While our selectivity profile for theenzyme reported under RefSeq accession no. WP_004927398 increased from octanal upto tetradecanal, we also found high activity for butanal and hexanal with this enzymeand found a decrease in the activity for hexadecanal (which was not tested as asubstrate for the FAldDH in that previous report [13]).

Of the six genes selected for this study, all but Maqu_0607 showed FAldDH activitywhen a range of different aldehyde substrates was tested. Efforts to obtain activity from

FIG 1 SDS-PAGE of various FAldDH enzymes from M. aquaeolei VT8 and A. baylyi following one-steppurification by metal affinity chromatography. The protein standard is shown on the left, followed by thesix enzymes, each with a primary band corresponding to a fatty aldehyde dehydrogenase. Labels includethe proposed protein apparent molecular mass based on the primary sequence from the geneticconstruct.

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Maqu_0607 were abandoned after multiple purifications failed to show any activitywith the substrates selected, and we instead focused our efforts on characterizing theremaining five enzymes that yielded FAldDH activity.

In addition to testing these enzymes with fatty aldehydes, we were also interestedin their activity toward acetaldehyde, as many of these enzymes still showed consid-erable activity toward substrates as small as butanal. Of the four enzymes from M.aquaeolei VT8, all showed some residual activity toward acetaldehyde, but Maqu_3410showed the greatest activity, which correlated with the broader enzyme activity foundfor this enzyme than those for the other three M. aquaeolei VT8 FAldDHs (Fig. 4). Theenzyme from A. baylyi reported under RefSeq accession no. WP_004927398 showed asignificantly higher level of activity with acetaldehyde than did any of the enzymesfrom M. aquaeolei VT8. This is in stark contrast to what was reported previously for theFAldDH from Acinetobacter sp. strain M-1, where those researchers did not report anyactivity with acetaldehyde (13).

We were also interested in testing aldehydes that share similarity to indigenous fattyacids. M. aquaeolei VT8 contains high levels of C16:1 and C18:1 fatty acids (3). However,

FIG 2 Specific activities of five fatty aldehyde dehydrogenase enzymes when assayed with a range of aldehyde substrates (C4 to C16) and residues that areproposed to align the aldehyde substrate binding site. The aldehyde substrates were incrementally increased in length from left to right (x axis). Assays wererun with 1.5 mM NAD� and 200 �M the indicated aldehyde, and the specific activity was monitored spectrophotometrically at 340 nm in a 1-cm-path-lengthcuvette with a 1-ml volume. Quantities of protein added ranged from 1 �g to 100 �g. Specific activity is reported as moles of NAD� reduced per minute permilligram of enzyme added (averages � standard deviations; n � 3), and reaction mixtures were maintained at 22°C. The alignment shows the regions of thevarious FAldDHs studied here or reported previously (13), with the residues proposed to line the aldehyde binding site highlighted. Residue numbering shownabove the alignment is based on the positions for the Maqu_3316-derived enzyme. The catalytic cysteine residue is Cys282 in this enzyme.

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the corresponding fatty aldehydes are not easily obtained through commercial sources,so we instead pursued a strategy to synthesize these fatty aldehydes from the corre-sponding fatty alcohol according to an oxidation protocol.

A comparison of the specificities of each of these enzymes with unsaturated versussaturated fatty aldehydes is shown in Fig. 5. Maqu_3410, Maqu_3572, and the enzymereported under RefSeq accession no. WP_004927398 all showed increased selectivitytoward C16:1 versus C16, while Maqu_0438 and Maqu_3316 showed comparable levelsof activity. All five enzymes were found to be active with C18:1 fatty aldehyde, althoughMaqu_3572 was the only enzyme that showed higher activity with C18:1 than with C16.In terms of the amino acid sequence alignments, Maqu_3410 and the enzyme reportedunder RefSeq accession no. WP_004927398 clustered together (Fig. 3) and also showedmore similar profiles with these potential native substrates containing unsaturatedbonds than with the straight-chain C16 fatty aldehyde (Fig. 5). Both enzymes showedthe highest specific activity with acetaldehyde of the five enzymes tested here, al-though the activity of the enzyme reported under RefSeq accession no. WP_004927398was significantly higher than those of all of the enzymes cloned from M. aquaeolei VT8(Fig. 4).

Enzyme selectivity for NAD� and NADP�. The oxidation of fatty aldehyde byFAldDH is dependent on the reduction of an electron-transporting cofactor. To deter-mine if these enzymes showed any differences in selectivity toward this electronacceptor, we tested each enzyme with both NAD� and NADP� as electron acceptors.

FIG 3 Phylogenetic tree of fatty aldehyde dehydrogenase enzymes examined in this study and six closelyrelated aldehyde dehydrogenase enzymes found in the Protein Data Bank. The PDB accession no.,GenBank accession no., or locus tag for M. aquaeolei VT8 is listed on the left, and the type of aldehydedehydrogenase and the host organism from which it was obtained are displayed on the right. Enzymesfrom the Protein Data Bank with the highest similarity to those of M. aquaeolei VT8 were selected forsequence comparisons (31, 32). The primary amino acid sequences were aligned and mapped phyloge-netically by using the software program Multalin (25).

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All five enzymes showed a higher specificity for NAD� than for NADP�, as would beexpected for catabolic processes (Fig. 6).

Optimal enzyme activity and temperature dependence. The selectivity assays(Fig. 2 to 6) were all performed at 22°C, while a previous study (13) of a bacterial FAldDHutilized a temperature of 43°C. To determine the enzyme stability and also themaximum activity that could be obtained, assays were performed by using the sub-strate with the highest activity (Fig. 2), and the enzymes were further tested by rampingup the temperature until activity was no longer linear throughout the assay. Table 1shows the relative activity at 22°C for the top substrate, the maximum activity that was

FIG 4 Specific activities of five aldehyde dehydrogenase enzymes when assayed with acetaldehyde.Assays were run with 1.5 mM NAD� and 200 �M acetaldehyde, and the specific activity was monitoredspectrophotometrically at 340 nm. Specific activity is reported as moles of NAD� reduced per minute permilligram of enzyme added (averages � standard deviations; n � 3), and reaction mixtures weremaintained at 22°C.

FIG 5 Comparison of substrate specificities of unsaturated aldehydes relative to the specific activitiesfound using the saturated aldehyde hexadecanal. Assays were performed with 1.5 mM NAD� and 200�M palmityl aldehyde (C16, saturated), palmitoleyl aldehyde (C16:1, unsaturated), or oleyl aldehyde (C18:1,unsaturated). The specific activity was measured spectrophotometrically at 340 nm, and reactions wereperformed at 22°C (averages � standard deviations; n � 3).

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obtained, and the temperature at which this maximum activity was obtained.Maqu_3572 showed the greatest temperature stability (53°C), with an �10-fold im-provement in activity versus what was found at 22°C for decanal. A similar improve-ment was found for the enzyme reported under RefSeq accession no. WP_004927398with butanal.

Overall structure and domain organization of the Maqu_3316 FAldDH. Basedon the purity and stability of specific FAldDH enzymes, we initiated crystallizationscreens and obtained two crystal structures for the Maqu_3316 FAldDH, one com-plexed with the substrate decanal and the other complexed with the cofactor NAD�.The two structures have been refined to 2.3-Å and 3.1-Å resolutions, respectively. Inboth structures, there are two FAldDH monomers, in the form of a symmetrical anddomain-swapped homodimer, existing in an asymmetric unit. Each monomer isL-shaped and consists of 487 amino acid residues, which are organized into threedomains: an N-terminal domain that binds the cofactor NAD� located at the pivot ofthe L shape (Fig. 7A, magenta); a catalytic domain at the short arm of the L shape withthe substrate decanal bound at the interface of the catalytic and N-terminal domains(Fig. 7A, ice blue); and, at the long arm of the L shape, a small oligomerization domain(Fig. 7A, green). The closest homologous structures currently available are thosereported under PDB accession no. 4KNA (N-succinylglutamate 5-semialdehyde dehy-drogenase from Burkholderia thailandensis) and 3JU8, with amino acid sequence iden-

TABLE 1 Activities assayed at elevated temperaturesa

Enzyme Substrateb

Avg activity (�mol ofNAD� reduced min�1

mg enzyme�1) � SDat 22°Cc

Avg highest activity (�molof NAD� reduced min�1

mg enzyme�1) � SD(temp [°C])c

Maqu_0438 C10 0.94 � 0.03 2.76 � 0.18 (47)Maqu_3316 C10 1.74 � 0.72 3.89 � 0.21 (43)Maqu_3410 C6 0.70 � 0.01 5.65 � 0.22 (45)Maqu_3572 C10 3.14 � 0.13 29.25 � 1.82 (53)RefSeq accession no.

WP_004927398C4 3.22 � 0.94 35.76 � 1.93 (47)

aReactions were performed at the highest temperature without activity loss, as described in Materials andMethods.

bEach enzyme was assayed with an aldehyde of the indicated length, chosen for substrates with the highestspecificity.

cn � 3.

FIG 6 Comparison of substrate specificities for NAD� relative to NADP�. Each enzyme was assayed with200 �M of the indicated aldehyde and 1.5 mM NAD� or NADP�. The specific activity was measuredspectrophotometrically at 340 nm, and reactions were performed at 22°C (averages � standard devia-tions; n � 3).

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tities of 63% and 62%, respectively. The root mean square deviations (RMSDs) forFAldDH with the homologs reported under PDB accession no. 4KNA and 3JU8 are 0.75and 0.66 Å, respectively, for all C� atoms (Fig. 7B).

FAldDH dimerization. The two monomers of FAldDH associate into a tightlyintertwined dimer arranged in a head-to-tail fashion (Fig. 8). The extensive interactionburies a total surface area of 3,950 Å2, with a formation energy of �28 kcal/molcalculated by using the program PISA (18). The interface interactions are mainlyhydrophobic: 559 nonbonded contacts compared to 10 salt bridges and 54 hydrogen-bonding interactions. The main architecture of the dimer is formed by the NAD�

binding domain and catalytic domain, with the extended oligomerization domainsdocking onto the groove of opposite monomers between the NAD� binding andcatalytic domains. The oligomerization domain contributes 3 �-strands to the �-sheetcore of the catalytic domain from the other molecule, stabilizing the FAldDH dimer.

NAD� binding pocket, active site, and substrate binding pocket. In the FAldDH/NAD� complex structure, each FAldDH monomer is bound with a copy of the NAD�

molecule. Similar to other homologous aldehyde dehydrogenase/NAD� complexes, theADP portion of NAD� has clear electron density, but the ribose sugar connecting thenicotinamide portion has very weak density. However, since the NAD� binding positionis known from previous structural studies of related aldehyde dehydrogenases (19) andthe nicotinamide has a clear density, intact NAD� was modeled. The extended NAD�

cofactor binding pocket is mainly hydrophobic. The ADP moiety of NAD� binds in apocket from the Rossmann fold N-terminal domain. The 2=-hydroxyl group is in closeproximity to C� of Ser175 (Fig. 9F), which could be relevant to the mechanism of theenzyme distinguishing between NAD� and NADP�. The nicotinamide has its ringstacked on the �S and C� atoms of the catalytic residue Cys282. A citrate binds to theputative active site and interacts with the �S atom of Cys282 through its terminalcarboxyl oxygen.

In the FAldDH/substrate complex, a continuous sausage-like electron density and aseparate, smaller, continuous electron density close to �S of Cys282 are observed. Dueto the discontinuity of the electron density, a molecule of ethylene glycol (EDO) ismodeled near �S of Cys282. The terminal oxygen atom of EDO pointing to �S of Cys282has a position very similar to that of the oxygen atom of citrate in the FAldDH/NAD�

complex. The longer sausage-like electron density was modeled with a nine-atomaliphatic chain, which represents the aliphatic tail of the substrate decanal, which wasused for soaking the crystal. Obtaining the complex of the aldehyde dehydrogenase

FIG 7 FAldDH monomer ribbon diagram. (A) Architecture of the three domains in a FAldDH monomer. The cofactor binding domain is in magenta, the catalyticdomain is in ice blue, and the oligomerization domain is in green. (B) Superposition of FAldDH onto monomers reported under PDB accession no. 4KNA and4JU8.

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with the linear substrate has been elusive despite a prolonged and intensive effort inthe field (20). Compared to the polyethylene glycol (PEG) molecule observed in thestructure reported under PDB accession no. 2VRO (Fig. 9B) (19), which was hypothe-sized to mimic the substrate tail, our results with the aldehyde decanal show that thetwo molecules have different positions: the PEG molecule in the structure reportedunder PDB accession no. 2VRO has one end facing the outside of the enzyme, while thedecanal in the present structure has both termini in the interior of the enzyme. Havingboth termini in the interior of the enzyme provides advantages for the enzymaticreaction: the interior of the enzyme could provide a more hydrophobic environment forthe substrate and could be more selective for the length of the substrate due to thepocket size limitation.

DISCUSSION

Fatty aldehydes pose a potential problem within the cell, as aldehydes are reactivemolecules that can be toxic at elevated concentrations. For organisms that producethese compounds as an intermediate in biosynthetic pathways, managing the levels ofinternal fatty aldehydes should be very important. For species that fill an importantenvironmental niche and utilize crude oil or other biologically derived lipids as agrowth substrate, the oxidation of these aldehydes would be essential for properfunction and meeting the energy needs of the cell. Fatty aldehydes have beenimplicated as a potential intermediate in the biosynthesis of fatty alcohols, althoughmany reports have shown that the enzymes in bacteria that reduce compounds fromthe fatty acid or activated fatty acid pools (acyl-CoA or acyl-ACP compounds) are alsohighly efficient at reducing fatty aldehydes and are not likely to release the fatty

FIG 8 Surface and ribbon representations of the dimerization of FAldDH monomers. The two monomers are shown in green and magenta, respectively. (A) Theethylene glycol bound in the active site and the bound decanal in the substrate binding pocket are shown in blue and dark brown, respectively (PDB accessionno. 5U0L). (B) The bound citrate in the active site and NAD� are shown in blue and dark brown sticks, respectively (PDB accession no. 5U0M). Abbreviations:EDO, ethylene glycol; CIT, citrate.

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aldehydes as a reaction intermediate (7, 9, 10, 21), while fatty aldehydes serve as aspecific intermediate in the biosynthesis of alkanes in cyanobacteria (16, 17). Interest-ingly, during a previous analysis of gene transcription in M. aquaeolei VT8 during theaccumulation of wax esters, it was found that one specific gene implicated in thepotential oxidation of fatty aldehydes (Maqu_3410) was highly transcribed during bothexponential growth and wax ester accumulation, even when utilizing citrate as agrowth substrate (8). This was somewhat surprising, since FAldDHs are generallyassumed to be associated with catabolic processes, and the production of wax estersis believed to be associated with energy storage, involving primarily anabolic processes(12, 13).

Ishige et al. (13) reported the characterization of a FAldDH obtained from Acineto-

FIG 9 Ligand binding and interactions in the binding pocket. (A) EDO and the fatty aldehyde substrate tail in the active site/substratepocket, with the 2mFo � DFc electron density map for the decanal tail contoured at 1.0 � depicted in blue mesh. (B) Ligands observedin the decanal/Maqu_3316 (ligands are decanal and EDO) complex and NAD�/Maqu_3316 (ligands are NAD� and CIT) complex structuresand in the structure reported under PDB accession no. 2VRO (the ligand is PEG) in the substrate binding pocket (the structures weresuperimposed to align the substrate binding pockets from each structure). (C) NAD� in the cofactor binding pocket, with the 2mFo � DFc

electron density map contoured at 1.0 � depicted in blue mesh. (D) The ligand EDO (blue) and the decanal tail (brown) in the substratebinding pocket and all the amino acids and water molecules (red balls labeled “W”) that are within 4 Å from the ligands. (E) Relativepositions of the cofactors, substrates, and substrate or product analogs in the substrate/cofactor binding pockets, with a 90° rotated viewof panel B around the vertical axis. (F) Cofactor NAD� interactions with the amino acids forming the cofactor binding pocket.

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bacter sp. strain M-1 that was able to utilize long-chain fatty aldehydes as a substrate.Since then, there have been few follow-up biochemical studies of these enzymes.Additionally, few reports have detailed the activity of these enzymes with substratesthat share similarity with potential native fatty-acid-derived substrates. A BLAST analysisof the genome of M. aquaeolei VT8 revealed quite a few genes that might code foradditional FAldDHs. In this work, we cloned and isolated five putative FAldDHs from M.aquaeolei VT8 and also cloned an additional FAldDH from A. baylyi. Of the six proteinsthat were obtained, only one (Maqu_0607) failed to show any activity with fattyaldehydes. We acknowledge that this lack of activity could be related to the instabilityof this enzyme during purification or a result of the incorporation of the polyhistidinetag to assist in purification, although the inclusion of the polyhistidine tag did not affectthe other five FAldDHs that were isolated and characterized. Several of the otherFAldDHs were also prone to large differences in activity for different purifications,although some activity could be rescued from many of these preparations by firstdegassing the protein preparation under an argon atmosphere and then adding a smallamount of �-mercaptoethanol. This approach was not successful for rescuing FAldDHactivity with Maqu_0607. Previous reports implicated potential cysteine residues aspossible sites that can become oxidized and inhibit enzyme activity (22). We alsoacknowledge the possibility that this enzyme might be active with a much morespecific substrate that was not tested as part of this study.

The genome of M. aquaeolei VT8 contains at least 10 different genes that code forenzymes with the potential to oxidize fatty aldehydes (15). Maqu_3410 (EC 1.2.1—) hadthe closest similarity score based on a BLAST search using the previously reported fattyaldehyde dehydrogenase from Acinetobacter sp. strain M-1 (13). It was followed closelyby another seven genes that had significant similarity scores (E values of �4e�38) andwere annotated as various aldehyde dehydrogenases. These included Maqu_0438 (EC1.2.1.3), Maqu_3572 (also annotated as a coniferyl-aldehyde dehydrogenase [EC1.2.1.68]), Maqu_3316 (also annotated as a succinylglutamate-semialdehyde dehydro-genase [EC 1.2.1.71]), and Maqu_0607 (EC 1.2.1.3). Each of these genes was cloned andpurified as part of this work. Additional genes not characterized included Maqu_3841(annotated as a betaine aldehyde dehydrogenase [EC 1.2.1.8]), Maqu_3647 (annotatedas a succinate semialdehyde dehydrogenase [EC 1.2.1.16]), and Maqu_2133 (annotatedas a methylmalonate-semialdehyde dehydrogenase [EC 1.2.1.18]). Several of thesegenes were also found to be highly transcribed during exponential growth and waxester accumulation, similar to what was found for Maqu_3410 (8), including Maqu_3316and Maqu_3572 (21).

Many of the substrates that were selected for this characterization were not easilyobtainable from commercial sources. For this reason, we instead chose to utilize asynthetic route to generate these substrates from commercially available fatty alcohols.Based on this work, we were able to characterize the substrate specificity of each ofthese enzymes with fatty aldehydes that can be derived from the indigenous fatty acidpool that has been characterized for M. aquaeolei VT8 in previous work (3, 8). Weacknowledge that the substrates selected in this study do not represent a comprehen-sive list of potential substrates (or even the most likely natural substrate, based onseveral of the annotations listed above), but these results provide a measure of activitywith small-, medium-, and long-chain fatty aldehydes of various lengths as well asseveral unsaturated fatty aldehydes. More importantly, all of the assays were performedby the same laboratory under the same conditions so that direct comparisons betweenthese different enzymes can be made with reasonable confidence. Several discretedifferences between the five FAldDHs that were isolated in an active form were found(Fig. 2, 4, and 5), and these differences correlate to some degree with the alignmentsshown in Fig. 3.

The enzyme corresponding to Maqu_3316 yielded diffraction-quality crystals. Thisenzyme showed a higher degree of specificity for a single substrate (decanal) than didany of the other enzymes characterized in this study with the selection of substratestested. We note that Maqu_3316 was also annotated as a potential succinylglutamate-

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semialdehyde dehydrogenase, and succinylglutamate-semialdehyde would have a sub-strate length similar to that of the substrate decanal, although it would have significantbranching groups and additional organic acid functional groups versus the simplestraight-chain aldehyde decanal.

The structure solved here with the decanal substrate bound to Maqu_3316 providesevidence of the fatty aldehyde hydrophobic substrate binding pocket for this enzyme.Based on the discontinuity of the electron density, we modeled a portion of thelong-chain aldehyde and a molecule of EDO to represent the electron density that wasobserved. Based on this density, we can define many of the residues that line thispotential fatty aldehyde substrate binding pocket in the Maqu_3316 enzyme. Sincethese residues are expected to mediate substrate selectivity, we performed a moredetailed analysis to show the general locations of these residues (Fig. 9D) and show thesequence conservation in the regions surrounding these specific residues from the fiveenzymes characterized in this study (Fig. 2, bottom) together with the differences insubstrate specificity with straight-chain aldehydes (Fig. 2, top). This analysis assumesthat each of these enzymes would share significant conservation of the enzyme foldand tertiary structure with the Maqu_3316-derived enzyme. However, strong foldconservation has been the case for this structure and other structures of relatedaldehyde dehydrogenase enzymes (Fig. 7B), so this should be a reasonable assumption.Based on these results, several specific residues of these enzymes are highlighted aspotential targets to alter the potential substrate specificity (Fig. 2). Most profoundly, theHis154, Leu155, Ser458, and Ala459 residues seem to be primary determinants ofsubstrate specificity. The Ser458 residue faces the C-8 position of decanal. InMaqu_0438 and Maqu_3572, Ser458 is replaced by a histidine residue, which couldinterfere with the binding of long-chain aldehydes, while Ala459 is replaced by aglycine residue, which should compensate for the histidine and provide more room andflexibility in the pocket, endowing the enzyme with more tolerance to shorter or longersubstrates. In Maqu_3410 and the enzyme reported under RefSeq accession no.WP_004927398, the Ser458 and Ala459 residues are replaced with asparagine andhistidine residues, respectively. These substitutions are larger and should create a morerigid binding site, such that shorter substrates would be preferred. Additionally, theHis154-Leu155 pair is replaced by leucine and methionine, respectively. This combina-tion may be more flexible, allowing for interactions with longer aliphatic aldehydes.These changes may result in the observed inverted bell-shaped specificity profiles forthese two enzymes. Studies to test this hypothesis could be pursued in the future todetermine whether mutations introduced into these sites result in changes of substratespecificity.

The two structures that were obtained in this study provide a view for how theFAldDH orients substrates to catalyze the oxidation of a fatty aldehyde to a fatty acid.These structures adopt a typical aldehyde dehydrogenase fold. In each of the twocomplexes, the substrate or cofactor contains a special “solvent” molecule bound in theactive site, which mimics the configuration of the substrate or the product in thecatalysis cycle. Figure 10A depicts the four-step enzymatic catalysis. In Fig. 10B, the twostructures are superimposed upon one another to bring all of the solvent/ligands intothe binding pockets near the active site of the enzyme. The complete decanal moleculeis modeled and refined in Fig. 10, and the �-carbon atoms of decanal, EDO, and CITeach overlap in this pocket, which is in close proximity to the catalytic S atom of Cys282.The �-carbon atom of the decanal depicts a longer distance from the S atom. EDO, asubstrate analog, occupies the active site, hindering the ability of the aldehyde to bindand blocking decanal access. The ethylene glycol forms two hydrogen bond interac-tions with the side-chain OH group of Thr438 (3.3 Å) and the S atom of Cys282 (2.2 Å),as shown in Fig. 9D. The C4 atom of NAD� lies 4.5 Å from the aldehyde oxygen atomof decanal. This distance is sufficient for the C4 atom of NAD� to accept the H atomfrom the �-carbon of the decanal during catalysis. Figure 10D shows a citrate moleculeoccupying the same site. The acid group of the citrate ligand is analogous to theterminal acid of the fatty acid product.

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Conclusions. The efforts undertaken in this study combined biochemical andstructural studies using a range of different substrates, including several that are notreadily available from commercial sources. The results obtained can be used to informthe rational selection of potential residues that might alter the selectivity of theseenzymes, which could be used in future biosynthetic approaches to tailor specificreactions. These results also reveal the potential for certain enzymes, such as thoseannotated as succinylglutamate-semialdehyde dehydrogenases, coniferyl-aldehyde de-hydrogenases, or benzaldehyde dehydrogenases, to perform an additional functionwithin the cell, as these enzymes cloned from M. aquaeolei VT8 all exhibited activitiessimilar to one another with a range of small-, medium-, and long-chain aldehydes.Future genetic studies could better identify the roles of these genes and their productsin the cell, while mutagenesis studies could probe the features of substrate specificity.

MATERIALS AND METHODSStrains and materials. Marinobacter aquaeolei VT8 (ATCC 700491) and Acinetobacter baylyi (ATCC

33305) were obtained from the American Type Culture Collection. Escherichia coli JM109 was obtainedfrom New England BioLabs (Ipswich, MA), while Escherichia coli BL21(DE3) was obtained from Novagen(Madison, WI). All chemicals and reagents were purchased from Sigma-Aldrich (St. Louis, MO) or FisherScientific (Pittsburgh, PA) unless otherwise specified. Restriction enzymes and T4 DNA ligase wereobtained from New England BioLabs. Coenzymes and fatty aldehydes were sourced from Sigma-Aldrich

FIG 10 Enzyme catalytic mechanism and atom configuration of the atoms of the substrate (decanal), the substrate analog EDO/FAldDH complex, and theproduct analog CIT/FAldDH complex. (A) Two-dimensional diagram of the four-step enzymatic catalysis performed by FAldDH. The substrate and productshown in red represent the two analogs in the present structures. (B) Superposition of the two complex structures showing all of the ligands in the cofactorand substrate binding pockets. The C4 atom of NAD� that participates in the reduction is labeled and shown in a stick representation. The Cys282 and Glu248residues that participate in the reaction are also shown. (C) The EDO ligand can be viewed as the aldehyde substrate analog and forms a hydrogen bond withthe S atom of Cys282 with a bond distance of 2.2 Å. (D) The CIT ligand with a carboxyl group can be viewed as the acid product analog, which reflects thelikely atomic configuration of the product/enzyme complex. The branched atoms are shown in high transparency to highlight the linear portion of the molecule.

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(St. Louis, MO) or Fisher Scientific (Pittsburgh, PA) or synthesized as follows. All PCRs were performed byusing the failsafe PCR enzyme system (Epicentre, Madison, WI).

Synthesis of fatty aldehydes. Long-chain fatty alcohols were obtained from Nu-Chek Prep (Elysian,MN), and a 100-�l volume of the fatty alcohol was dissolved in methylene chloride (0.1 M) with 10 mol%2,2,6,6-tetramethylpiperidyl-1-oxyl (TEMPO) and 1:1 equivalents of phenyl iodonium diacetate (PIDA).Aliquots were removed to monitor the progress of the reaction by thin-layer chromatography (TLC) using4% ethyl acetate (EtOAc) in hexanes (Hex) as the eluent and visualized by using a KMnO4 stain. Thesolution was mixed overnight at room temperature in a sealed round-bottom flask. The sample was thenanalyzed by TLC to determine if it had reached the endpoint, and the volatiles were removed by rotatoryevaporation at 35°C under reduced pressure. The sample was then placed under a vacuum in alaboratory HVAC system to remove any remaining volatile material. The mixture of products was thenanalyzed by using TLC (4% EtOAc in Hex) to obtain Rf values of the products. The product was purifiedby column chromatography (silica gel, 4% EtOAc in Hex) using vacuum pressure to purify the aldehyde,and the isolated fraction was again dried by rotary evaporation and an HVAC system. The aldehyde wasdissolved in CDCl3 and analyzed by 1H nuclear magnetic resonance (NMR) spectroscopy to characterizethe desired product and assay purity. For tetradecanal (C14H28O), 1H NMR (400 MHz, CDCl3) � 9.76 (t, J �1.9, 1H), 2.42 (td, J � 7.1, 1.9, 2H), 1.63 (p, J � 7.3, 2H), 1.36 to 1.20 (m, 20H), 0.88 (t, J � 6.8, 3H). Forhexadecanal (C16H32O), 1H NMR (500 MHz, CDCl3) � 9.77 (t, J � 1.9, 1H), 2.42 (td, J � 7.4, 1.9, 2H), 1.63(p, J � 7.4, 2H), 1.37 to 1.19 (m, 24H), 0.88 (t, J � 7.0, 3H). For palmitoleyl aldehyde (C16H30O), 1H NMR(400 MHz, CDCl3) � 9.76 (t, J � 1.9, 1H), 5.40 to 5.29 (m, 2H), 2.42 (td, J � 7.4, 1.9, 2H), 2.01 (q, J � 6.2,4H), 1.63 (p, 7.2, 2H), 1.40 to 1.20 (m, 16H), 0.88 (t, J � 6.7, 3H). For oleyl aldehyde (C18H34O), 1H NMR (400MHz, CDCl3) � 9.76 (t, J � 1.9, 1H), 5.40 to 5.29 (m, 2H), 2.42 (td, J � 7.4, 1.9, 2H), 2.01 (q, J � 6.0, 4H),1.63 (p, J � 7.2, 2H), 1.38 to 1.20 (m, 20H), 0.88 (t, J � 6.7, 3H). Once these aldehydes were obtained, theywere stored frozen under an argon atmosphere until required for testing.

Plasmid constructions. The gene for a putative fatty aldehyde dehydrogenase (Maqu_3410) from M.aquaeolei VT8 was cloned by PCR with primers BBP801 (5=-GACTACCATGGAATCTATGCACAACCCGGACAAGAAGGCTCCGTTG-3=) and BBP802 (5=-GTCATTCTAGAATCAGAAGAACCCGAGAGGGTTGGTGTCGTAGCTG-3=) and then digested with the restriction enzymes NcoI and XbaI (underlined in primer sequencesfor clarity) and ligated into plasmid pBB114, a pUC19 derivative containing kanamycin in place ofampicillin resistance (8). Following sequencing to confirm the gene, this DNA segment was shuttled,using the same restriction enzymes, into a pET-19b derivative plasmid to create plasmid pETMFA, whichincludes an 8� polyhistidine tag at the N terminus.

The gene for a second putative fatty aldehyde dehydrogenase (Maqu_0438) from M. aquaeolei VT8was inadvertently cloned by PCR with primers BBP1778 (5=-NNNGGATCCATGGACAGATTGCTAGTCTGGCGGAGGTTGG-3=) and BBP1779 (5=-NNNAAGCTTCAGCCCAGAATACGAACAGCAGCATCTATG-3=) and thendigested with the restriction enzymes BamHI and HindIII and ligated into plasmid pBB114. Two mistakesfound in the primer regions of this construct were repaired by site-specific mutagenesis (Stratagenemethod; Agilent Technologies, Santa Clara, CA). Following sequencing to confirm the gene, this DNAsegment was shuttled, using NcoI and HindIII, into a pET-19b derivative plasmid to create plasmidpPCRMFA9, which includes an 8� polyhistidine tag at the N terminus.

The gene for a third putative fatty aldehyde dehydrogenase (Maqu_0607) from M. aquaeolei VT8 wasalso cloned by PCR with primers BBP1778 and BBP1779 and then digested with the restriction enzymesBamHI and HindIII and ligated into plasmid pBB114. Site-specific mutagenesis was used to insert a silentmutation that removed a second NcoI site within the gene. Following sequencing to confirm the gene,this DNA segment was shuttled, using NcoI and HindIII, into a pET-19b derivative plasmid to createplasmid pPCRMFA10, which includes an 8� polyhistidine tag at the N terminus.

The gene for a fourth putative fatty aldehyde dehydrogenase (Maqu_3572) from M. aquaeolei VT8was cloned by PCR with primers BBP2319 (5=-NNNAAGCTTCAGCGAATAAACAGCTTATACACCAACC-3=)and BBP2320 (5=-NNNATGCATCACCACCATCATCACGGTGCCACCGTCGTCCAGCTCACC-3=) and then di-gested with the restriction enzymes NsiI and HindIII and ligated into plasmid pBB114. Followingsequencing to confirm the gene, this DNA segment was shuttled, using NdeI and HindIII, into a pET-19bderivative plasmid to create plasmid pPCRMFA36, which includes a 6� polyhistidine tag at the Nterminus.

The gene for a fifth putative fatty aldehyde dehydrogenase (Maqu_3316) from M. aquaeolei VT8 wascloned by PCR with primers BBP2343 (5=-NNNAAGCTTGGTTGGCCACCGCATTGACGTTGG-3=) and BBP2344(5=NNNTCTAGAGTAACCACCATACCCATGAACTGCATCG-3=) and then digested with the restriction en-zymes HindIII and XbaI and ligated into plasmid pBB114. A polyhistidine tag was then added to this geneby using primers BBP2420 (5=-NNNGGTACCATATGCATCACCACCATCACCATGCAAACCTGACAGGCAATGTGTACATC-3=) and BBP2421 (5=-GATCCCCGGGTACCGAG CTCGAATTCACTG-3=) and then digesting theproduct with KpnI and ligation. Following sequencing to confirm the gene, this DNA segment wasshuttled, using NdeI and HindIII, into a pET-19b derivative plasmid to create plasmid pPCRMFA47, whichincludes a 6� polyhistidine tag at the N terminus.

The gene for a sixth putative fatty aldehyde dehydrogenase (RefSeq accession no. WP_004927398)from Acinetobacter baylyi was cloned by PCR with primers BBP2389 (5=-NNNATGCATCACCACCATCATCACCGTTATATCGATCCTAATCAACCTGGCTC-3=) and BBP2390 (5=-NNNAAGCTTAGAAGAAGCCCATTGGTTTTGTTGAATAAC-3=) and then digested with the restriction enzymes NsiI and HindIII and ligated into plasmidpBB114. Following sequencing to confirm the gene, this DNA segment was shuttled, using NdeI andHindIII, into a pET-19b derivative plasmid to create plasmid pPCRMFA48, which includes a 6� polyhis-tidine tag at the N terminus.

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Protein purification. Completed plasmids were transformed into E. coli BL21(DE3) for the expressionof the protein. Cells were grown in Miller’s lysogeny broth (LB) at 30°C and induced with isopropyl-�-D-thiogalactopyranoside (IPTG) (50 mg liter�1) for 2 h, when they reached an optical density at 600 nmof 0.6. Cells were harvested by centrifugation at 7,000 � g for 7 min. Protein was extracted in a mannersimilar to what was described previously, using metal affinity chromatography followed by a desaltingcolumn for equilibration in a final buffer containing 50 mM potassium phosphate (pH 7.0) and 300 mMNaCl (23). Final isolated fractions were pooled and tested for purity by running the fractions on SDS-PAGEgels. The protein was either flash frozen in liquid nitrogen or stored at 4°C.

Fatty aldehyde dehydrogenase activity assays. Enzyme activity assays were conducted withquartz cuvettes with a 1-cm path length using a Varian Bio 50 UV-visible (UV-Vis) spectrophotometerequipped with temperature control. Each assay mixture contained 900 �l of reaction buffer (25 mMglycine and 50 mM potassium phosphate [pH 9.6]). Aldehydes were diluted in isopropanol such that 200nmol was added to the cuvette. NAD� was then added from a concentrated stock solution to bring thefinal concentration to 1.5 �mol. The various components were mixed thoroughly with a pipette beforebeing placed into the spectrophotometer. Samples were monitored by measuring the absorbance at 340nm for 1 min to confirm a stable baseline before the addition of the enzyme (10 to 100 �g) to initiatethe reaction. The production of NADH was determined by correlating the absorbance change to theextinction coefficient for NADH (6,220 M�1 cm�1 at 340 nm). Initial rates of reaction were calculated byexporting the raw data to Microsoft Excel (Microsoft, Redmond, WA) to convert the absorbance perminute to specific activity rates. NADP�-based assays were performed according to the same procedurebut with NADP� substituted for NAD�. All calculations are based on data from a minimum of 3 replicates(n � 3). Saturated aliphatic aldehyde substrates tested included acetaldehyde (C2), butanal (C4), hexanal(C6), octanal (C8), decanal (C10), dodecanal (C12), tetradecanal (C14), and hexadecanal (C16). Unsaturatedaliphatic aldehyde substrates tested included palmitoleyl aldehyde (C16:1) and oleyl aldehyde (C18:1).Quantification of enzyme concentrations used for calculating specific activities were based on theabsorbance of the protein at 280 nm and the extinction coefficient calculated with the ExPaSy ProteinParameters algorithm (see http://web.expasy.org/protparam/) (24).

TABLE 2 Data collection and refinement statisticsa

Parameter

Value(s) for complex:

FAldDH/substrate FAldDH/NAD�

Data collection statisticsResolution range (Å) 92.04–2.29 (2.37–2.29) 78.33–3.08 (3.29–3.08)Space group P41212 P41212Unit cellb (Å) 98.71, 98.71, 254.73 99.38, 99.38, 254.60Total no. of reflections 841,693 (79,325) 178,295 (31,311)No. of unique reflections 57,590 (4,945) 24,631 (4,349)Multiplicity 14.6 (14.3) 7.2 (7.2)Completeness (%) 97.41 (86.95) 99.90 (99.90)I/�I 14.82 (0.84) 6.30 (0.70)Rmerge 0.176 (4.78) 0.371 (2.64)Rmeas 0.182 (4.96) 0.399 (2.84)Rp.i.m. 0.04713 (1.282) 0.1446 (1.035)CC1/2 0.999 (0.563) 0.991 (0.505)

Refinement statisticsNo. of reflections 56,699 (4,937) 23,468 (2,126)No. of reflections for Rfree 2,742 (235) 1,139 (98)Rwork (%) 22.62 (42.82) 21.78 (40.43)Rfree (%) 26.72 (46.56) 27.30 (43.39)No. of nonhydrogen atoms 7,402 7,475

Macromolecules 7,350 7,350Ligands 4 92Solvent 48 33

No. of protein residues 975 975RMSD

Bond length (Å) 0.002 0.005Bond angle (°) 0.61 0.72

Ramachandran plot (%)Favored regions 96.1 95.9Allowed regions 3.9 4.1Outliers 0 0

Avg B factor (Å2) 73.92 86.09Macromolecules 73.98 85.84Ligands 88.34 121.80Solvent 62.38 43.41

aStatistics for the highest-resolution shell are shown in parentheses.bValues shown are for dimensions a, b, and c.

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Sequence comparisons. Enzymes similar to the original fatty aldehyde dehydrogenase from Acin-etobacter sp. strain M-1 (13) and the genes cloned here were used to perform a BLAST search. Enzymesthat shared high identity and had three-dimensional (3D) models in the Protein Data Bank werecompared to our selection of enzymes. The sequences of the enzyme reported under RefSeq accessionno. WP_004927398, Maqu_3316, Maqu_0438, Maqu_3572, Maqu_3410, and six closely related sequenceswere compared by using Multalin (http://multalin.toulouse.inra.fr/multalin/) (25). Sequence alignmentswere generated along with a hierarchical tree of similarity.

Crystallization and structure determination of a FAldDH. Purified Maqu_3316 at 36 mg/ml wasscreened for crystallization by the sitting-drop vapor diffusion method at the Nanoliter CrystallizationFacility at the Department of Biochemistry, Molecular Biology and Biophysics, University of Minnesota,using the CrystalTrak system (Rigaku, Tokyo, Japan). In total, 10 screens, each consisting of 96 uniqueconditions, were set up. Crystals appeared after 3 weeks under multiple conditions. Crystals weretransferred into cryoprotectant solutions containing the corresponding reservoir solution supplementedwith 20% (vol/vol) ethylene glycol. For ligand soaking, the cryoprotectant solution contained 20 mMdecanal or NAD�. The typical soaking time was 1 to 2 h. The crystals were then flash frozen in liquidnitrogen.

Crystals were screened at the Advanced Photon Source Northeastern Collaborative Access Teambeamlines (24-ID-C and 24-ID-E). The best-diffracting crystals were the ones from the MCSG_2 screen(Anatrace, Maumee, OH) (well ID F8, 0.2 M NH4H2PO4, 50% [vol/vol] 2-methyl-2,4-pentanediol, 0.1 M Tris[pH 8.5]) and the PEGRx HT screen (Hampton Research, Aliso Viejo, CA) (well ID F10 containing 0.2 Mammonium citrate, 20% [wt/vol] PEG 2,000 monomethyl ether, 0.1 M imidazole [pH 7.0]). The collecteddata were processed with XDS (26). The Matthews coefficient calculation indicated that there would likelybe a dimer of Maqu_3316 in an asymmetric unit. Using the monomer of succinylglutamic semialdehydedehydrogenase from Pseudomonas aeruginosa (PDB accession no. 3JU8) as a search model, molecularreplacement by PHASER (27) located two copies of the FAldDH monomers in the asymmetric unit.Subsequent iterative refinement with PHENIX suite (28) and model inspection and building using COOT(29) resulted in final Rwork/Rfree values of 22.62% and 26.72% for the FAldDH/substrate complex and21.78% and 27.30% for the FAldDH/NAD� complex, respectively. A summary of the data collection andrefinement statistics is shown in Table 2. Ramachandran analysis shows that 96.1%, 3.9%, and 0% of theprotein residues are in the most favored, allowed, and disallowed regions for the FAldDH/substratecomplex and that 95.9%, 4.1%, and 0% of the protein residues are in the most favored, allowed, anddisallowed regions for the FAldDH/NAD� complex, respectively. Molecular graphic images were pro-duced by using PYMOL (http://www.pymol.org/).

Accession number(s). The structure factors and coordinates have been deposited in the ProteinData Bank (30) under accession no. 5U0L and 5U0M, respectively.

ACKNOWLEDGMENTSWe thank Nagendra Palani for assistance in constructing plasmid pETMFA. We thank

Chris Rothstein and Zeyuan Wu for assistance in early characterization of theMaqu_3410 enzyme.

This work was supported by grants from the National Science Foundation to B.M.B.(award no. 0968781 and CBET-1437758) and C.J.D. (CHE-1151547) and from the Na-tional Institutes of Health to H.A. (NIGMS R35-GM118047). Further support was pro-vided to B.M.B. through generous startup funds through the University of Minnesota.This work is based upon research conducted at the Northeastern Collaborative AccessTeam beamlines, which are funded by the U.S. National Institutes of Health (NIGMSP41-GM103403). The Pilatus 6M detector on the 24-ID-C beamline is funded by anNIH-ORIP HEI grant (S10 RR029205). This research used resources of the AdvancedPhoton Source, a U.S. Department of Energy (DOE) Office of Science User Facilityoperated for the DOE Office of Science by Argonne National Laboratory under contractno. DE-AC02-06CH11357.

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