Biochimica et Biophysica Acta 1844 (2014) 681–693

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Aldehyde dehydrogenase enzyme ALDH3H1 from Arabidopsis thaliana: Identification of amino acid residues critical for cofactor specificity Naim Stiti, Karolina Podgórska, Dorothea Bartels ⁎ Institute of Molecular Physiology and Biotechnology of Plants, University of Bonn, Kirschallee 1, 53115 Bonn, Germany

a r t i c l e

i n f o

Article history: Received 12 June 2013 Received in revised form 9 January 2014 Accepted 13 January 2014 Available online 23 January 2014 Keywords: Aldehyde Cofactor Dehydrogenase activity NAD+ NADP+

a b s t r a c t The cofactor-binding site of the NAD+-dependent Arabidopsis thaliana aldehyde dehydrogenase ALDH3H1 was analyzed to understand structural features determining cofactor-specificity. Homology modeling and mutant analysis elucidated important amino acid residues. Glu149 occupies a central position in the cofactor-binding cleft, and its carboxylate group coordinates the 2′- and 3′-hydroxyl groups of the adenosyl ribose ring of NAD+ and repels the 2′-phosphate moiety of NADP+. If Glu149 is mutated to Gln, Asp, Asn or Thr the binding of NAD+ is altered and rendered the enzyme capable of using NADP+. This change is attributed to a weaker steric hindrance and elimination of the electrostatic repulsion force of the 2′-phosphate of NADP+. Simultaneous mutations of Glu149 and Ile200, which is situated opposite of the cofactor binding cleft, improved the enzyme efficiency with NADP+. The double mutant ALDH3H1Glu149Thr/Ile200Val showed a good catalysis with NADP+. Subsequently a triple mutation was generated by replacing Val178 by Arg in order to create a “closed” cofactor binding site. The cofactor specificity was shifted even further in favor of NADP+, as the mutant ALDH3H1E149T/V178R/I200V uses NADP+ with almost 7-fold higher catalytic efficiency compared to NAD+. Our experiments suggest that residues occupying positions equivalent to 149, 178 and 200 constitute a group of amino acids in the ALDH3H1 protein determining cofactor affinity. © 2014 Elsevier B.V. All rights reserved.

1. Introduction Aldehyde dehydrogenases (ALDHs): (E.C 1.2.1.3) constitute a large evolutionary conserved superfamily of enzymes that catalyze the irreversible oxidation of a wide range of aldehydes to their corresponding carboxylic acids using NAD(P)+ as cofactors. Aldehydes are widespread molecules that take part in different physiological processes. Aldehydes become toxic when they accumulate to excessive, nonphysiological levels. Hence, ALDHs are considered to be detoxifying enzymes which maintain physiological levels of aldehydes [1]. ALDHs are ubiquitous and are found throughout prokaryotic and eukaryotic organisms. With the completion of several genome sequencing projects, the number of newly identified ALDH genes from different taxa is steadily increasing, providing new insights into sequence plasticity and evolution [2–9]. Each ALDH monomeric subunit consists of three domains: the catalytic domain, the NAD(P)+-binding domain and the oligomerization domain. Subunits are assembled by hydrogen bonds into enzymatically active homotetramers for members of families 1, 2, 7, and 9 [10–13], whereas members of family3 function as homodimers [14,15].

Abbreviations: ALDH, aldehyde dehydrogenase; AMADH, aminoaldehyde dehydrogenase; BADH, betaine aldehyde dehydrogenase; ROS, reactive oxygen species ⁎ Corresponding author. Tel.: +49 228 73 2070; fax: +49 228 73 1697. E-mail address: [email protected] (D. Bartels). 1570-9639/$ – see front matter © 2014 Elsevier B.V. All rights reserved. http://dx.doi.org/10.1016/j.bbapap.2014.01.008

Plant ALDHs have been classified into 13 distinct protein families, where seven families (ALDH2, ALDH3, ALDH5, ALDH6, ALDH7, ALDH11 and ALDH18) have mammalian orthologs, and six families (ALDH10, ALDH12, ALDH21, ALDH22, ALDH23 and ALDH24) are specific for plants. Analysis of the genome sequence of Arabidopsis thaliana revealed 16 ALDH genes encoding members of ten ALDH protein families [3]. Plant ALDHs are localized in different subcellular compartments including cytosol, mitochondria, plastids, peroxisomes and microsomes [16–23]. This differential localization indicates functional specialization. Different cell compartments require appropriate ALDH enzymes with distinct physico-chemical characteristics. Developmental-dependent and organ-specific expression patterns of ALDHs have been reported [20,22–25]. The role of plant ALDHs in different physiological processes has become an emerging theme in plant physiology during the last years. The first plant ALDH to which a function was assigned was the spinach ALDH10 (betaine aldehyde dehydrogenase) which is involved in the synthesis of the osmoprotectant glycine betaine [26]. Later, the maize mitochondrial ALDH2B2/RF2A gene was identified as a nuclear restorer of cytoplasmic male sterility [27,28]. The mitochondrial class 2 ALDH of rice was reported to be essential for rapid detoxification of acetaldehyde which is produced upon re-aeration after submergence [29]. Expression of members of plant ALDHs from families 2, 3, 5, 7 and 10 is responsive to oxidative and abiotic stresses [25,26,29–31]. The involvement of these genes in stress protection has been demonstrated

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by analyzing transgenic plants. Overexpression of Arabidopsis ALDHs of families 3 and 7 confers tolerance to heavy metal, osmotic and oxidative stresses [21,32]. Likewise, ectopic expression of a soybean ALDH7 enhanced stress tolerance [33]. Involvement of ALDHs in stress tolerance was further corroborated by the analysis of Arabidopsis ALDH3I1, ALDH7B4 T-DNA and rice OsALDH7 knockout mutants which exhibited higher stress sensitivity [21,25]. Although physiological data imply an important function of plant ALDHs in growth and environmental stress adaptation, the structural properties of plant ALDHs remain mostly unveiled [25,34–36]. Solved crystal structures of ALDHs [10,11,14,15,37–39] revealed common structural features. Dehydrogenases bind nicotinamide cofactors in a region containing a core Rossmann-type fold domain, which consists of six parallel β-strands interspersed by α-helices that appear on both sides of the six-stranded β-strand. A typical glycine-rich motif (Gly1-X-Gly2-X-X-Gly3) forms a tight turn between the end of the first β-strand (β1) and the beginning of the so-called “dinucleotide binding helix” (αA) [40]. Structural variations have been found for the Rossmann fold in dehydrogenases. The Rossmann fold of ALDHs is different from other dehydrogenases. It is made up of a five stranded open α/β domain. Moreover, the NAD + -binding fingerprint (Gly 1 -X-Gly 2 -X-X-Gly 3 ) which forms a tight turn and is usually present between β1 and αA is missing. Consequently, the loop located between β1 and αA is more extended and bulkier. The classical tight turn is found between β4 and αD in ALDHs and corresponds to the characteristic glycine-rich sequence Gly1-X-X-X-XGly2 involved in interaction with the cofactor nicotinamide ring. The NAD(P)+ binding cleft is positioned between the helices αC and αD instead of the central helices as in alcohol dehydrogenases [14]. The relevance of different residues from the five open α/β strands of the Rossmann fold in cofactor recognition and accommodation was studied using site-directed mutagenesis approaches [34,41–43]. The residue located immediately downstream of the β2-strand functions as part of the NAD+ recognition site, since the 2′-hydroxyl group of the ribose moiety forms a hydrogen bond with its side chain [11,14]. The mutation of this residue changed the binding and the catalytic efficiency of the nicotinamide cofactor. Substitution of the equivalent residue of the rat ALDH3a1 in either Glu140 (Uniprot: P11883) by Asn, Gln or Thr favored NADP+ binding as shown by catalytic efficiency [41]. On the other hand, mutating the equivalent residue, Thr175 in the NADP+dependent Vibrio harveyi ALDH (VIBHA_ALDH Uniprot: Q56694) to Glu or Gln strengthened interaction with NAD+ [42]. The mutation of the adjacent Lys (Lys137 in ALDH3a1) to Glu affected cofactor-binding abilities negatively [41]. Similar observations were made when the equivalent residue in human liver mitochondrial ALDH2 was mutated (K192Q) [43]. In contrast, mutation of the last residue of the glycine rich sequence, Gly1−X-X-X-X-Gly2 in ALDH3a1, (Gly192) did not affect enzyme efficiency [41]. Very little is known about biochemical characteristics of families 2 and 3 plant ALDHs. Most biochemical studies in plants consist of kinetic analyses where substrate specificities were investigated taking into account the aliphatic chain length or their degree of unsaturation [34,35]. Arabidopsis family 3 ALDHs (ALDH3H1 and ALDH3I1) were found to be able to oxidize medium- to long-chain aliphatic aldehydes, with a preference for long-chain aldehydes. Affinities were also found to be generally lower with unsaturated than saturated aldehydes [34]. Crystal structures of ALDHs from different organisms were solved in their apo or holo forms, but there are very few reports on structural data of plant ALDHs. Homology-based structural reconstitutions of rice and maize ALDHs have been reported [44,45]. Recently, the first crystal structure of a chloroplastic ALDH10 from spinach SoBADH (Uniprot: 17202, PDB: 4A0M) was solved and amino acid residues critical for high BADH activity and substrate affinity have been identified [46]. Information is scarce about amino acid residues which are involved in modulating plant ALDH activities or which are critical in nicotinamide cofactor recognition and binding. We recently identified residues

involved in redox regulation of the family3 stress-responsive A. thaliana ALDHs, ALDH3H1 and ALDH3I1 and evidence was provided that Ile200 is a key residue responsible for the NAD+-dependency in A. thaliana ALDH3H1 [34]. In this study, we continued our investigation in cofactor binding modalities in the A. thaliana ALDH3H1 enzyme based on knowledge of cofactor specificities in other ALDHs, sequence comparisons, and site-directed mutagenesis. We engineered an active NADP+-specific enzyme, where substitutions were introduced in the cofactor binding pocket. Alteration of nicotinamide cofactor specificity provided new insights into plant family 3 ALDHs, as it identified amino acids which determine nicotinamide cofactor binding sites. 2. Results 2.1. Sequence alignment and prediction of amino acid residues involved in cofactor binding sites Recently we have demonstrated that the amino acid residue Ile200 is responsible for the Arabidopsis ALDH3H1 NAD+-dependency [34]. Ile200 is specific for ALDH3H1 and is replaced by Val in enzymes which are able to use NAD+ or Gly in the NADP+-dependent ALDH of V. harveyi VIBHA_ALDH, while it is present in the NADP+-dependent glyceraldehyde-3-phosphate dehydrogenase from Streptococcus mutans GAPN_STRMU (Fig. 1). We had indications that besides Ile200 also other amino acids contribute to the cofactor interaction. Therefore sequence alignments and cofactor binding site structure comparisons were performed to identify amino acids which are additionally involved in the ALDH3H1 cofactor binding. Amino acid sequences encompassing the cofactor binding site were aligned from the following enzymes: the NADP+-preferring ALDHs Rattus norvegicus ALDH3a1 (UniProt: P11883), Homo sapiens ALDH3A1 (UniProt: P30838), the NADP+-dependent S. mutans glyceraldehyde3-phosphate dehydrogenase (UniProt: Q59931), NADP+-specific fatty aldehyde dehydrogenase from V. harveyi (UniProt: Q56694), NAD+dependent A. thaliana ALDH3H1 (UniProt: Q70DU8), NAD+-preferring A. thaliana ALDH3I1 (UniProt: Q8W033) and the NAD+-dependent aldehyde dehydrogenase from Rhizobium meliloti (UniProt: Q930S8). These alignments revealed the presence of a Thr residue in the NADP+dependent dehydrogenases in a position where Glu is present in ALDHs which use NAD+ (Fig. 1) and site-directed mutagenesis experiments showed that the Thr residue contributes to the NADP+-dependency [38,42]. Secondary and tertiary structural models of Arabidopsis ALDH3H1 were built using crystal structures of rat ALDH3a1 (PDB: 1AD3) and human ALDH3A1 (PDB: 3SZB) as templates (Fig. 2). The stereochemical quality of the predicted protein structure was evaluated by analyzing residue-by-residue geometry and overall structure geometry using tools as ANOLEA energy [47], GROMOS force field energy [48] provided by SWISS-MODEL server and PROCHECK [49] which attest favorable energy environment showing the absence of misfolding of the whole enzyme and the absence of abnormal packing energy or aberrant geometries for amino acids of the cofactor binding site. Residues from 10 to 90 form five helices α1 (10–25), α2 (27–26), α3 (32–60), α4 (64–69) and α5 (73–90). The strand β0 (105–112) forms an arm-like stretch which bridges the α5 helix and the cofactor binding domain. The open α/β nicotinamide cofactor binding Rossmann fold consists of the five β-strands β1 (115–119), β2 (142–146), β3 (170–174), β4 (191–195) and β5 (215–218) which are interspersed by four α-helices αA (127–138), αB (152–165), αC (178–186) and αD (198–211) (Fig. 2A). The glycine-rich motif Gly1-X-X-X-X-Gly2, positioned between β4 and αD and characteristic for ALDHs corresponds to the sequence GSSKIG (residues 196–201). The catalytic region is made up of six parallel β-strands β6 (224–227), β7 (258–261), β9 (332–334), β10 (352–358), β11 (376–380), β12 (398–401), the antiparallel strand β8 (315–318) and six helices α10 (233–244), α11

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Fig. 1. Alignment of amino acid sequences encompassing cofactor-binding sites from different dehydrogenases. Amino acid sequences correspond to Arabidopsis thaliana (ARATH), ALDH3H1 (UniProt: Q70DU8) and ALDH3I1 (UniProt: Q8W033), Rattus norvegicus ALDH3a1 (UniProt: P11883), Vibrio harveyi VIBHA_ALDH (UniProt: Q56694), Homo sapiens HOMO_ALDH3A1 (UniProt: P30838), the NAD+-dependent Sinorhizobium meliloti RHIME_ALDH (UniProt: Q930S8), and the NADP+-dependent glyceraldehyde-3-phosphate dehydrogenase from Streptococcus mutans GAPN_STRMU (UniProt: Q59931). Alignment was performed using the ClustalW web-based multiple sequence alignment tool. Conserved residues are shaded in black. Residues that are important for NAD+/NADP+ preference are indicated in colors. Glu and Val which are conserved in class 3 ALDHs are indicated in green. The unusual residues (Thr, Ile, and Gly) are shaded in red or pink. Residues targeted for mutagenesis are indicated by downward-pointing arrows. Numbers in blue on the left-hand and right-hand side for each sequence indicate the positions of the amino acids in the native proteins.

(263–281), α12 (285–287), α13 (297–307), α14 (361–370), α15 (384–393). Like in rat ALDH3a1 and human ALDH3A1, the C-terminal polypeptide chain of Arabidopsis ALDH3H1 extends over the Rossmann fold forming α17 (429–435) and β13 (436–443). The oligomerization domain mediating the formation of the hydrogen-bonded active homodimers corresponds to the strands β12 and β13 as well as the helix αD (Fig. 2B). In the model the nicotinamide cofactor molecule, NAD+, is docked to the cofactor binding cleft and its binding properties were compared with other holoenzymes. This confirmed the important positions occupied by Glu149 and Ile200 supporting a role in cofactor interactions and the catalytic process (Fig. 3). The Glu149 side-chain carboxyl oxygen atoms lie at hydrogen-bonding distance to the adenine ribose of NAD+ in family 3 ALDHs tertiary structures, while Ile200 is situated at the opposite side of Glu149 and it is suggested to be important in adjusting the width of the cofactor-binding cleft. 2.2. Site-directed mutagenesis and kinetic analysis Based on the results above, Glu149 and Ile200 were chosen as targets for site-directed mutagenesis in the Arabidopsis ALDH3H1 enzyme. It was

tested whether mutations of Glu149 and Ile200 would alter enzyme specificity toward cofactors and would thus permit to elucidate their roles in cofactor recognition and binding. To examine the role of Glu149, it was mutated to Asp, in which the acidic residue is more distant from the ribose moiety of the NAD+. Glu149 was also changed to Gln to delete the acidic group or Asn and Thr to simultaneously remove the carboxylic acid group and to enlarge the cofactor binding cleft. The mutation of Glu149 was combined with mutations of Ile200 to amino acids with shorter side chains, i.e., Val or Gly. All mutant enzymes were expressed as recombinant proteins, purified and used for enzyme assays. The substrate specificity was determined by plotting the enzymatic activities against increasing hexanal concentrations at saturated cofactor concentrations. Two mM hexanal was found to be the optimal substrate concentration for all mutated enzymes and it was used in the subsequent tests for cofactor studies (Fig. A.2). All data are presented in Table 1. Single and double mutations at positions 149 and/or 200 have marked effects on cofactor affinities. The mutated enzymes showed a disruption in the interaction with NAD+, reflected by an increase of the KmNAD+ value and a decrease of catalytic efficiency kcatNAD+/ KmNAD+. The opposite effect is seen on NADP+ specificity and the related

Fig. 2. Predicted secondary and tertiary structure. (A) Predicted secondary structure components: arrows represent β-strands and ribbons represent α-helices. Helices and strands are annotated according to the convention used for alcohol dehydrogenases. Components highlighted in green correspond to the Rossmann fold (five strands connected by four helices). The glycine-rich motif, Gly1-X-X-X-X-Gly2 [14] (residues 196 – 201) and the active site cysteine, Cys253 [34] are denoted by orange and red boxes respectively. (B) Ribbon diagram of the predicted three-dimensional structure of A. thaliana ALDH3H1. The molecule is colored in gray. The cofactor-binding Rossmann-fold domain is depicted in green. The catalytic cysteine residue is shown in red. The predicted structure was built using the web-based modeling server SWISS-MODEL [53] and the Rattus norvegicus ALDH3a1 (PDB: 1AD3) as well as the human ALDH3A1 (PDB: 3SZB) as templates. Regions corresponding to helix αD, strands β12 and β13 (underlined with dotted red lines) are potentially involved in hydrogen bond-mediated oligomerization resulting in the functional homodimer as observed for ALDH3a1 [14].

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Fig. 3. Illustration of interactions of adenine ribose moieties with amino acid residues in the cofactor-binding site of different dehydrogenases. Location of NAD+, NADP+ or NAP2 (ligand homologeous to NADP+) adenine ribose moieties in the cofactor-binding site of the previously solved structures of dehydrogenases of ALDH3a1 from Rattus novegicus, (PDB: 1AD3), Vibrio harveyi VIBHA_ALDH, (PDB: 1EYY), HOMO_ALDH3A1 from Homo sapiens (PDB: 3SZA), RHIME_ALDH from Sinorhizobium meliloti (PDB: 3U4J), GAPN_STRMU from Streptococcus mutans (PDB: 2EUH) and the predicted model of the A. thaliana ALDH3H1. Cofactor-binding sites are magnified from the ribbon diagram of the crystal structure of ALDH3a1 (panel A), VIBHA_ALDH (panel B) ALDH3A1 (panel C), ALDH3H1 (panel D), RHIME_ALDH (panel E) monomers and the predicted model of ALDH3H1. Amino acid residues assumed to be important for the cofactor-binding are presented in stick model and highlighted in violet for Glu, blue for Thr, red for Ile, orange for Val, purple for Gly. Cofactors NAD+ (panels A,C,D,E), NADP+ (panel F) and ligand NAP-2 (homologeous to NADP+) (panel B) are shown in stick model and atoms are depicted as follows: oxygen, red; carbon, green; phosphorus, orange; nitrogen, blue; hydrogen atoms are hidden. The predicted structure of ALDH3H1 was generated by the Web-based server SWISS-MODEL [53], and rendered using PyMol.

catalytic efficiency: KmNADP+ decreased to reach a lower value than KmNAD+, whereas the catalytic efficiency with NADP+ (kcatNADP+/KmNADP+) increased. The substitution of Glu149 by Gln slightly affects the width of the cofactor binding cleft and resulted in the removal of a negative charge in vicinity of positions 2′ and 3′ of the ribose moiety of the cofactor. The corresponding mutated enzyme exhibits a loss of specificity for NAD+, as seen by the 2-fold increase of the KmNAD+ value (886 μM versus 421 μM for the wild-type enzyme) and a decrease of the catalytic

efficiency by one-fourth (12.7 s− 1 versus 17.2 s− 1 for the wild-type enzyme). The mutated enzyme also acquired the ability to use NADP+, although with a high KmNADP+ (2281 μM). On the basis of the catalytic efficiency, the ALDH3H1E149Q mutant enzyme still shows a greater preference for NAD+ than NADP+ as indicated by the specific catalytic efficiency ratio, (kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+) of 9.2, which is considered as a measure of specificity for NAD+ relative to NADP+. The mutated enzyme ALDH3H1E149D has an anion-side chain in position 149 which is one methyl group shorter compared to the wild-type

Table 1 Kinetic properties of recombinant ALDH3H1 and mutant enzymes. Apparent Km and kcat values were determined for NAD+ and NADP+ with hexanal as substrate at saturating concentrations. Catalytic efficiency is expressed as kcat/Km(app) (s−1) and characterizes aldehyde-oxidizing capacity. Results are mean values ± S.E.M. from at least three independent experiments. Enzymatic characteristics related to ALDH3H1 were reported previously [34]. Enzyme

ALDH3H1 ALDH3H1E149N ALDH3H1E149Q ALDH3H1E149D ALDH3H1E149T ALDH3H1I200V ALDH3H1E149N/I200V ALDH3H1E149Q/I200V ALDH3H1E149D/I200V ALDH3H1E149T/I200V ALDH3H1I200G ALDH3H1E149N/I200G ALDH3H1E149Q/I200G ALDH3H1E149D/I200G ALDH3H1E149T/I200G

NAD+

NADP+

(kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+)

KmNAD+ (μM)

kcatNAD+ (s−1)

kcatNAD+/KmNAD+ (s−1.mM−1)

KmNADP+ (μM)

kcatNADP+ (s−1)

kcatNADP+/KmNADP+ (s−1.mM−1)

421 ± 23.5 630 ± 33.1 886 ± 110 441 ± 40 1262 ± 38 496 ± 4 992 ± 54.4 1856 ± 43 991 ± 82 1649 ± 38 3218 ± 54 4136 ± 124 4059 ± 130 3651 ± 125 3418 ± 187

17.2 ± 1.6 16.6 ± 0.7 12.7 ± 0.8 10.7 ± 0.8 10.2 ± 0.4 18.5 ± 1.7 13.1 ± 0.6 11.4 ± 0.6 12.4 ± 0.4 15.7 ± 0.3 16.3 ± 1.7 26.9 ± 2.1 20.1 ± 1.9 23.2 ± 0.6 33.9 ± 0.8

40.9 ± 1.2 26.3 ± 0.9 14.3 ± 0.7 24.3 ± 1.1 8.1 ± 0.2 37.3 ± 1.1 13.2 ± 0.5 6.1 ± 0.1 12.5 ± 0.6 9.5 ± 0.7 5.1 ± 0.3 6.5 ± 0.2 5 ± 0.4 6.4 ± 0.3 9.9 ± 0.6

n.a 1335 ± 82 2281 ± 106 3461 ± 249 1294 ± 56 2300 ± 76 1176 ± 36 1225 ± 24 1046 ± 175 675 ± 12 1817 ± 17 3212 ± 42.7 1877 ± 43.7 2635 ± 167.6 2148 ± 67.2

n.a 2.6 ± 0.04 3.5 ± 0.09 1.7 ± 0.04 7.4 ± 0.2 1 ± 0.09 5.3 ± 0.2 6.1 ± 0.2 4.4 ± 0.9 5.1 ± 0.4 0.6 ± 0.09 2.3 ± 0.09 2.3 ± 0.04 1.3 ± 0.06 1.1 ± 0,02

n.a 1.96 ± 0.03 1.56 ± 0.04 0.49 ± 0.09 5.68 ± 0.1 0.45 ± 0.05 4.49 ± 0.2 5 ± 0.16 4.21 ± 0.8 7.54 ± 0.6 0.31 ± 0.06 0.71 ± 0.08 1.21 ± 0.04 0.48 ± 0.02 0.48 ± 0.03

n.a 13.4 9.2 49.6 1.4 83.4 2.9 1.2 3 1.3 16.4 9.2 4.1 13.1 20.7

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enzyme. This mutation only had a small effect on NAD+-interaction with a K mNAD + value of 441 μM versus 421 μM for the wild-type enzyme but the corresponding catalytic efficiency was reduced by 40%. ALDH3H1E149D is able to use NADP+ with a high KmNADP+ of 3461 μM and a low catalytic efficiency (kcatNADP+/KmNADP+ = 0.49 s−1.mM−1). In the mutated enzyme ALDH3H1E149N, the values of KmNAD+ and KmNADP+ changed to 630 μM and 1335 μM respectively upon removal of the negative charge. The catalytic efficiency for NAD+ decreased by one-third compared to the wild-type enzyme (26.3 s−1.mM−1 for ALDH3H1E149N vs. 40.9 s− 1.mM− 1 for the wild-type ALDH3H1), although the enzyme still shows a higher efficiency with NAD + ((kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+) = 13.47). More pronounced alterations of the cofactor specificity occurred upon replacement of Glu149 by Thr. ALDH3H1E149T showed similar Km values for both cofactors coupled with a 5-fold drop in the catalytic efficiency for NAD+ (kcatNAD+/KmNAD+ = 8.1 s−1.mM−1 vs. 40.9 s−1.mM−1 for the wild-type ALDH3H1). This yielded an enzyme which can use NAD+ and NADP+ with a decrease of the specific catalytic efficiency ratio to reach 1.42. These observations provide support for the hypothesis that the Thr residue caused the change of nicotinamide cofactor specificity from NAD+-dependency and the increase of related aldehyde dehydrogenase activity. We also investigated kinetic parameters of enzymes carrying two mutations, where the width of the cofactor binding cleft varies concomitantly with different electrostatic charges in the vicinity of the adenine ribose of NAD(P)+ (Table 1). Compared to the non-mutated and all tested single mutated enzymes, the double mutant ALDH3H1E149Q/I200V showed the strongest decrease in terms of NAD+ binding (KmNAD+ = 1856 μM) and retained only 15% of its catalytic efficiency with NAD+, while recognition and binding of NADP+ improved. The value KmNADP+ of 1225 μM was for the first time lower than KmNAD+ (KmNAD+ = 1856 μM). This double mutant enzyme is able to use both cofactors with equal effectiveness as shown by the ratio of specific catalytic efficiencies (k catNAD + /K mNAD + )/(k catNADP + /K mNADP + ) = 1.2. The observed dual cofactor specificity is caused by the low turnover number k catNADP + value (k catNADP + = 6.1 s − 1 ) when compared to kcatNAD+ (kcatNAD+ = 11.4 s− 1). The KmNAD+ for the double mutant ALDH3H1E149D/I200V showed a nearly two-fold increase compared to the wild-type enzyme (K mNAD + = 991 μM vs. 421 μM respectively) and a K mNADP + of 1046 μM was reached. This double mutation lowered the NAD+ catalytic efficiency k catNAD + /K mNAD + by 70% while k catNADP + /K mNADP + reached the value 4.21 s − 1 .mM − 1 but was still 3 times lower than the catalytic efficiency with NAD + (12.5 s − 1 .mM − 1 ). A very similar effect on kinetic parameters was observed for the double mutant ALDH3H1 E149N/I200V . The amino acid substitutions weakened the NAD + -binding, as seen from the K mNAD + value of 992 μM, which remained about two times higher than the wild-

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type enzyme. The KmNADP+ reached the same range as observed for ALDH3H1E149D/I200V (KmNADP+ = 1176 μM). Similarly to the double mutant ALDH3H1 E149D/I200V ALDH3H1 E149N/I200V prefers NAD + as shown by the comparison of specific catalytic efficiencies for each cofactor, despite that Km values are almost in the same range. The mutated enzyme carrying the double mutation E149T/ I200V altered the preference for cofactor binding and it performed well with both cofactors compared to the wild-type enzyme. ALDH3H1 E149T/I200V displayed a weaker binding of NAD + and an improved recognition of NADP + as shown by its 2.5-fold lower KmNADP+ compared to K+ mNAD (675 μM versus 1648 μM respectively). ALDH3H1 E149T/I200V is quite unique among all tested enzymes in that the K mNADP + is the lowest and the catalytic efficiency with NAD+ is 4-fold lower than the wild-type enzyme, while the catalytic efficiency with NADP+ is increased. On the basis of the specific catalytic efficiency ratio (kcatNAD+ /K mNAD +)/(k catNADP+ /KmNADP +) of 1.3, the enzyme shifted from an NAD+-dependency to dual cofactor specificity and it is almost equally effective with both cofactors, NAD+ and NADP+. This improved ability to use NADP+ was also observed when besides hexanal other substrates (trans-2-hexenal, nonanal and trans-2nonenal) were oxidized by ALDH3H1E149T/I200V at saturation concentrations (Fig. A.2). The change of substrate does not alter the NADP+binding ability and its preference as cofactor (Table 2A). The lowest value for KmNADP+ was observed for the oxidation of trans-2-hexenal (168 μM). KmNADP+ values are lower when unsaturated aliphatic aldehydes were used compared to their saturated homologs. The catalytic efficiency (kcatNAD+/KmNAD+) decreased by about 10-fold compared to the wild-type enzyme when trans-2-hexenal and trans-2-nonenal were used (Tables 1 and 2). All observed specific catalytic efficiency ratios (kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+) correspond to dual cofactor specificity. The lowest ratio (0.71) corresponds to the dehydrogenation of trans-2-nonenal, which indicates a shift toward NADP+-specificity. The Ile200 to Gly mutation (mutated isoforms: ALDH3H1I200G, ALDH3H1 E149Q/I200G , ALDH3H1 E149D/I200G , ALDH3H1 E149N/I200G , ALDH3H1 E149T/I200G ) impaired affinities for both cofactors as shown by high KmNAD+ and KmNADP+ values (Table 1). Although maximal activities are higher, especially for the double-mutant forms, it cannot compensate the loss of cofactor binding ability (high Km values), thus the catalytic efficiencies for NAD+ were reduced and dropped to less than 15% for some mutants compared to the wild-type enzyme ALDH3H1. It was tested whether the affinity toward NADP+ could be further improved with additional mutations. Therefore, Val178 was mutated to Arg in the double mutant ALDH3H1E149T/I200V to yield the mutant ALDH3H1E149T/V178R/I200V. This mutation shifted the cofactor preference from NAD+ toward NADP+. On the basis of the catalytic efficiency ratio (k catNAD + /K mNAD +)/(k catNADP+ /KmNADP +), the triple mutant, ALDH3H1E149T/V178R/I200V, displays a 7-fold stronger preference

Table 2 Kinetic properties of the recombinant double mutant enzyme ALDH3H1E149T/I200V (A) and the triple mutant ALDH3H1E149T/V178R/I200V (B). Apparent Km and kcat values were determined for NAD+ and NADP+ with different aldehyde substrates at saturating concentrations. Catalytic efficiency is expressed as kcat/Km(app) (s−1) and characterizes aldehyde-oxidizing capacity. Results are mean values ± S.E.M. from at least three independent experiments. NAD+

NADP+

(kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+)

KmNAD+ (μM)

kcatNAD+ (s−1)

kcatNAD+/KmNAD+ (s−1.mM−1)

KmNADP+ (μM)

A Hexanal trans-2-Hexenal Nonanal trans-2-Nonenal

1648 ± 38 1092 ± 42 1829 ± 61 564 ± 27

15.7 ± 0.3 3.1 ± 0.1 18 ± 0.3 2.2 ± 0.1

9.5 ± 0.2 2.8 ± 0.1 9.9 ± 0.5 4 ± 0.2

675 ± 18 168 ± 2 470 ± 18 292 ± 3

5.1 ± 0.4 0.4 ± 0.03 3.5 ± 0.1 1.2 ± 0.1

7.5 ± 0.5 2.2 ± 0.2 7.6 ± 0.4 5.6 ± 0.5

1.26 1.27 1.30 0.71

B Hexanal trans-2-Hexenal Nonanal trans-2-Nonenal

2524 ± 70 1622 ± 112 2115 ± 15 1068 ± 31

12.2 ± 0.8 1.8 ± 0.02 12.7 ± 0.3 4 ± 0.04

4.8 ± 0.2 1.1 ± 0.1 6 ± 0.2 3.8 ± 0.1

277 ± 12 103 ± 2 222 ± 1 91 ± 1.4

9 ± 0.05 1.8 ± 0.03 13.3 ± 0.9 2.7 ± 0.1

32.9 ± 1.4 18 ± 0.1 59.6 ± 4.4 29.8 ± 0.5

0.15 0.06 0.10 0.13

kcatNADP+ (s−1)

kcatNADP+/KmNADP+ (s−1.mM−1)

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for NADP+ over NAD+ with hexanal as substrate (Table 2B). The observed change in cofactor specificity is explained by the increase of KmNAD+ accompanied by a small change in kcatNAD+. Consequently, the enzymatic efficiency with NAD+ (kcatNAD+/KmNAD+) also decreased 8-fold when compared to the wild-type enzyme. These changes of enzyme performance related to KmNAD+ as cofactor occurred simultaneously with a decrease of KmNAD+ value and almost a 2-fold increase of the kcatNADP+ compared to the double mutant ALDH3H1E149T/I200V. The acquired ability to use NADP+ reached a similar level in terms of catalytic efficiency when compared to the initial performance with NAD+, which was observed for the wild-type ALDH3H1 (kcatNAD+/KmNAD+ = 40.9 s−1.mM−1 for ALDH3H1 versus kcatNADP+/KmNADP+ = 32.9 s−1.mM−1 for the triple mutant ALDH3H1E149T/V178R/I200V). The amplitude of the shift in cofactor specificity from NAD+ to NADP+ in the triple mutant depends on the oxidized substrate. Based on the catalytic efficiency ratio ((kcatNAD+/KmNAD+)/(kcatNADP+/KmNADP+) =0.15), the triple mutant ALDH3H1E149T/V178R/I200V displays nearly a 6-fold preference for NADP+ over NAD+ when hexanal was used. Besides hexanal other substrates were oxidized at saturation concentrations (Fig. A.2). The catalytic efficiency ratio decreased to 0.06 with trans-2-hexenal as substrate (Table 2B). This corresponds to an increase of NADP + preference over NAD + by a factor of 2.5 when the substrate hexanal was substituted by its unsaturated conjugate trans-2-hexenal. The use of trans-2-hexenal as substrate is also correlated to a decrease of KmNADP+ which reaches 103 μM, and corresponds to a more efficient NADP + binding by a factor of three when compared to hexanal (KmNADP+ = 277 μM) (Table 2B). The improved NADP+ binding with unsaturated substrates was also observed when trans-2-nonenal was oxidized. With this substrate the KmNADP+ showed a three-fold decrease when compared to nonanal (91 μM versus 222 μM respectively) (Table 2B). In constrast to aliphatic C6 aldehydes (hexanal and trans-2-hexenal), the change of the unsaturation degree in C9 aldehydes (nonanal and trans-2-nonenal) does not affect the nicotinamide cofactor preference and the catalytic efficiency ratio did not change (0.10 for nonanal and 0.13 for trans-2-nonenal) (Table 2B). Binding studies were performed to determine the equilibrium dissociation constants relative to both cofactors (KdNAD + and KdNADP+) for ALDH3H1 wild-type enzyme as well as the triple mutant ALDH3H1E149T/V178R/I200V in the presence of NADH and NADPH. These studies were done to confirm the observations deduced from the calculation of kinetic parameters (see above, KmNAD+, KdNADP+, kcatNAD+, kcatNADP+ and caluculated catalytic efficiencies). The Michaelis–Menten and Lineweaver–Burk plots show that NADH and NADPH are competitive inhibitors of NAD+ and NADP+ (Fig. 4). For the Lineweaver–Burk plot, the intercept 1/V (1/ALDH activity) remained unchanged and only the slope varied with increasing concentrations of NADH (Fig. 4B). The Michaelis–Menten plots show that the maximum rate of the reaction (Vmax) is not altered after adding the reduced forms of nicotinamide cofactors at different concentrations (Fig. 4A, C, D, E and F). The affinity of the wild-type enzyme decreased 2.5-fold upon addition of 250 μM of NADH. Dissociation constant value for NAD+, KdNAD+ was 920 ± 49 μM in the presence of 250 μM of NADH. KdNAD+ values are correlated with the concentrations of the added NADH. KdNAD+ values were 489 ± 19 μM and 1132 ± 49 μM when the dehydrogenase reaction was inhibited using 100 μM or 375 μM of NADH respectively (Fig. 4A). Two hundred-and-fifty μM of NADH have almost no effect on the dehydrogenase activity for ALDH3H1E149T/V178R/I200V, since the Michaelis– Menten saturation curves are nearly similar in the absence or presence of 250 μM NADH (Fig. 4B and C) and the corresponding KdNAD+ (2857 ± 78 μM) and KdNADP+ (362 ± 3 μM) differ only slightly from KmNAD+ (2524 ± 70 μM) and KmNADP+ (277 ± 12 μM). To confirm the improvement of NADP+ binding of the triple mutant ALDH3H1E149T/V178R/I200V, KdNAD+ and KdNADP+ were determined in the presence of different concentrations of NADPH. In contrast to NADH,

the effect of NADPH was stronger by preventing NAD+ to bind to ALDH3H1E149T/V178R/I200V. The saturation plateau was not reached even at high concentrations of NAD+ (from 5 mM to 10 mM) in the presence of the same concentration (250 μM) or even lower concentrations of NADPH (20 μM, 50 μM, and 100 μM) (Fig. 4E). The saturation plateau was reached when the concentration of NADPH was reduced to 10 μM and the corresponding dissociation constant K dNAD + (2400 ± 98 μM) represents an increase of about 600 μM when compared to the determined KmNAD+ for this enzyme without adding the reduced form of the coenzyme for the same test (Fig. 4E). NADPH also affected the affinity of the enzyme to NADP+. The saturation plateau was not reached when NADPH was added at concentrations of 250 μM or 375 μM, but it was reached when the concentration of NADPH was decreased to 100 μM (Fig. 4F). The corresponding dissociation constant value KdNADP+ (515 ± 14 μM) was found to be 2-fold higher than KmNADP+ which reflects a weakening of the NADP+ binding. The effect of the triple mutation on substrate specificity is negligible. Similarly to the wild-type enzyme, ALDH3H1E149T/V178R/I200V prefers longer chain aliphatic aldehydes when NAD+ is used at saturating concentrations (nonanal and trans-2-nonenal over hexanal and trans-2-hexenal respectively) although the catalytic efficiency for the latter substrates is lower than for the wild-type ALDH3H1 (Table 3). Catalytic efficiency showed that similar to the wild-type enzyme, ALDH3H1E149T/V178R/I200V prefers saturated aliphatic aldehydes over their corresponding unsaturated conjugates. Based on catalytic efficiency values, under saturating concentrations of NAD+ shorter saturated aliphatic aldehydes (C6 aldehydes) are preferred like it was observed for the wild-type ALDH3H1 (Table 3). 3. Discussion Understanding the structural features which determine cofactorspecificity is of particular interest in the enzymology of ALDHs. In our study, we used a systematic screening approach to identify the molecular determinants of cofactor-specificity. We performed a multiple sequence alignment and a comparison of cofactor-binding cleft structures with selected ALDH sequences to identify important amino acid residues. These residues were targeted by site-directed mutagenesis to identify the molecular parameters that control cofactor-specificity. 3.1. ALDH3H1 enzymes with a mutation at position 149 Cofactor-specificity in family3 ALDHs is determined by several amino acids, but most importantly by the Glu residue located at position 149, which interacts with the adenine ribose or the 2′-phosphate of NAD+ and NADP+ respectively. Glu149 occupies a central position in the cofactor-binding cavity and coordinates the adenine ribose hydroxyl groups of NAD+, while the negative charge of its side chain repels the 2′phosphate moiety of NADP+ accounting for the increased KmNADP+ value for most family3 ALDHs when compared to KmNAD+ [41]. Furthermore, the glutamyl residue that has been thought to be incompatible with the binding of the 2′-phosphate of NADP+ and therefore promoting the NAD+-dependence in ALDHs is present in Pseudomonas aeruginosa BADH (PaBADH, Uniprot: Q9HTJ1, PDB: 2VE5) but the enzyme is still able to use NADP+. This is because the electrostatic repulsion force exerted from the glutamyl residue on the 2′-phosphate group of NADP+ is abolished. The side chain of Glu179 is directed away from the 2′-phosphate by anionic interaction with Arg40 which abolishes the electrostatic repulsion causing a rejection of NADP+ as cofactor [50]. The presence of the Glu residue at position 149 in ALDH3H1 is not the only factor discriminating against NADP+. The Glu residue is exerting an electrostatic repulsion against the 2′-phosphate moiety of the NADP+. The width of the cofactor binding cleft is a critical parameter which influences the accommodation of NADP+. In fact, the

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Fig. 4. Effect of NAD(P)H on ALDH3H1 and ALDH3H1E149T/V178R/I200V inactivation and binding affinities in the presence of NAD(P)+. Michaelis–Menten and Lineweaver–Burk plots illustrating that ALDH3H1 affinity toward NAD+ depends on NADH concentrations (panels A & B) Michaelis–Menten plots illustrating that ALDH3H1E149T/V178R/I200V affinities toward NAD+ and NADP+ are not affected by NADH concentrations (panels C & D). Michaelis–Menten plots illustrating that ALDH3H1E149T/V178R/I200V affinities to NAD+ and NADP+ depend on the NADPH concentrations (panels E & F).

Table 3 Kinetic properties related to substrate specificity of wild-type enzyme ALDH3H1 and triple mutant enzyme ALDH3H1E149T/V178R/I200V. Apparent Km and kcat values were determined for different substrates at high of concentration of NAD+ (4 mM for the triple mutant which correspond to the saturation for ALDH3H1). Catalytic efficiency is expressed as kcat/Km(app) (s−1) and characterizes aldehyde-oxidizing capacity. Results are mean values ± S.E.M. from at least three independent experiments. Enzymatic characteristics related to ALDH3H1 were reported previously [34]. Substrate

Hexanal trans-2-Hexenal Nonanal trans-2-Nonenal

ALDH3H1

ALDH3H1E149T/V178R/I200V

Km(μM)

kcat (s−1)

kcat/Km (s−1.mM−1)

Km (μM)

kcat (s−1)

kcat/Km (s−1.mM−1)

71 ± 12 180 ± 24 8±2 3 ± 0.7

11.2 ± 0.5 2.2 ± 0.1 17.9 ± 3.8 2.7 ± 0.17

154 ± 3 12.4 ± 0.8 2166 ± 37 918 ± 21

72 ± 3.4 199 ± 6 65 ± 4.9 32.2 ± 1.8

8.2 ± 0.1 1.1 ± 0.04 13 ± 0.5 3.8 ± 0.1

115 ± 5 5.3 ± 0.2 213 ± 27 120 ± 7

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chloroplastic isoform, ALDH3I1, has a Glu residue in this position (Glu212) but it is still able to use NADP+ as cofactor. This is due to a larger cofactor binding cleft resulting from the presence of Val instead of Ile (Ile200) (equivalent to a loss of a methyl group) at the opposite side of the cofactor binding cavity when compared with ALDH3H1. Probably, for the same reason, the mutants ALDH3H1Ile200Val and ALDH3H1Ile200Gly were able to use NADP+ while the electrostatic repulsing force resulting from E149 is still effective [34]. In the case of the chloroplastic isoform ALDH3I1, the reverse mutation V263I failed to convert the enzyme from an NAD(P)+ into an NAD+-dependent enzyme [34]. Because the length of the side chain of the amino acid residue located opposite of the glutamyl residue in the cofactor binding cleft cooperates with other amino acids yet to be identified are important in binding and stabilizing the cofactor. Our investigation focused on the Glu 149 residue in the Arabidopsis ALDH3H1 enzyme. Substitution of the Glu149 residue by an uncharged residue, i.e., Gln or Asn, rendered the enzyme able to use NADP+ as cofactor and simultaneously reduced the catalytic efficiency with NAD+ as illustrated by higher KmNAD+ and lower kcatNAD+/KmNAD+ values than the non-mutated enzyme. In the mutant ALDH3H1E149Q the electrostatic repulsion force neighboring the adenosyl ribose ring of the nicotinamide cofactor has been abolished. Gln offers a side chain nearly of the same length as the charged glutamate, thus its insertion at position 149 is not expected to change the width of the cofactor-binding cleft. The only difference is that its uncharged amide group does not repel the 2′-phosphate group of NADP+. In addition, its amide nitrogen can form a hydrogen-bond involving the terminal oxygen atoms of the 2′-phosphate of the adenosine ribose. These changes in terms of electrostatic forces are the primary reasons for the observed disruption of NAD+-binding and the acquired ability to use NADP+ (Table 1). The presence of Gln149 does not change the distance between the side chain and the adenine ribose of NAD+, but it alters the ability of hydrogen-bonding. While the original Glu carboxyl side chain interacts with the ribose moiety via forming two hydrogen-bonds involving the 2′- and 3′-hydroxyl groups of the adenosyl ribose ring of the NAD+, the Gln amide oxygen side chain can only form one. Thus the physicochemical properties of the side chain of the Gln149 can explain the weaker binding of NAD+ as shown by the two-fold increase of KmNAD+ value and the two-third decreased catalytic efficiency compared to the nonmutated enzyme (Table 1). The amide side chain of Asn presents the same potential as Gln to interact with the 2′-phosphate group of NADP+. Thus, substitution of Glu by Asn also yielded an enzyme (ALDH3H1E149N) capable of using NADP+. Acceptance of NADP+ as cofactor can be attributed to the concomitant effect of weakening the steric hindrance and elimination of the electrostatic repulsion force in the region surrounding the 2′-phosphate group of the adenine moiety. Compared to ALDH3H1E149Q, the shorter side chain appears to promote the NADP+-binding, as deduced from a nearly 2-fold decrease of the KmNADP+ value and the one-fourth higher catalytic efficiency. The removal of a methyl group may provide an additional space for a better accommodation of the 2′-phosphate group and thus allowing the NADP+ molecule to interact with the appropriate distance of the residue in position 149. On the other hand, as observed for ALDH3H1E149Q, the presence of an amide oxygen side chain instead of a carboxyl side chain weakened the NAD+-binding when compared to the mutant ALDH3H1E149D showing the equivalent acidic residue in the same position (Table 1). The aspartic acid in the mutant ALDH3H1E149D has one methyl group less and thus reduces the carboxylate anionic repelling side chain and consequently allowed the adenosine ribose to shift. Thus the 2′-phosphate moiety of the ribose is closer to the cofactor-binding pocket. This promotes tighter binding and better interaction with NADP+. This shows that the space required to accommodate the phosphate group is not sufficient in the native NAD+-dependent enzyme due to the presence of the bulky Glu149.

The displacement of the negatively charged carboxylate group also weakened the hydrogen-bonding ability and thus negatively affected the interaction with the 2′- and 3′-hydroxyl groups of the ribose moiety of NAD+. This led to a slightly higher KmNAD+ and a lower catalytic efficiency (62% of the initial efficiency) compared to the wild-type enzyme (Table 1). A similar situation where ALDH3H1 acquired the ability to use NADP+ was observed when the glutamate carboxylate anionic repelling side chain was conserved in the region surrounding the 2′-phosphate group and a smaller side chain amino acid was positioned opposite to the cleft at position 200 (mutation I200V) [34]. This shows that widening the cofactor binding cleft by the size of one methyl group opposite to the ribose moiety is crucial for the ability to use NADP+ when a repulsive electrostatic force is present at position 149. Kinetic characteristics of ALDH3H1E149T showed an improved performance in terms of using NADP+ as cofactor. Evidence for the change in cofactor specificity is derived from the relative degree of NADP+ binding (KmNADP+/KmNAD+ = 1.02), which is 2- to 8-times lower than that observed for other mutants. An increase of the turnover number kcatNADP+ is observed conversely to that relative to NAD+. This resulted in a rise of the catalytic efficiency with NADP+ and a concomitant decrease for NAD+. This finding shows that the Thr149 residue is involved in cofactor recognition and shifted cofactor-specificity. The Thr149 promoted the NADP+-binding due to its small size and the restricted dihedral angles of its β-methyl group which creates a cavity accommodating the 2′-phosphate of the ribose of adenosine in NADP+. Thr149 forms hydrogen bonds with the phosphate group and its side chain hydroxyl group as well as the main chain amide. The effect of Thr149 with regards to the enhancement of NADP+-binding is consistent with observations from crystallographic structures of the NADP + -dependent glyceraldehyde-3-phosphate dehydrogenase from S. mutans (GAPN_STRMU) [39] and the fatty aldehyde dehydrogenase from the luminescent bacterium V. harveyi (VIBHA_ALDH) [51] where Thr residues (Thr180 and Thr175 respectively) occupy similar positions in the cofactor-binding cleft (Figs. 1 and 3). These residues coordinate NADP+ via hydrogen bonds between their hydroxyl groups and the terminal oxygen atoms of the 2′-phosphate group. The crucial role of Thr occupying such a position in the cofactor-binding site was confirmed by substitution of Thr175 of the fatty aldehyde dehydrogenase of V. harveyi by the negatively charged residues Asp or Glu, which changed the cofactor-specificity from NADP+ to NAD+. The replacement of Thr by the polar Gln generated a more efficient NAD+-specific enzyme without loss of NADP+-dependent activity [42]. 3.2. ALDH3H1 enzymes with double mutations at positions 149 and 200 3.2.1. Double mutant enzymes carrying a Val residue at position 200 Simultaneous replacement of Glu149 by Thr and Ile200 by Val resulted in the most efficient enzyme with NADP+ as cofactor so far when hexanal is used as substrate (Table 1). Changing the substrate to a longer chain aldehyde or corresponding unsaturated aldehydes confirmed the kinetic properties of this double mutant enzyme. The Michaelis constants showed that changing the substrate had little effect on the cofactor preference, since the ratios (kcatNAD+/KmNAD+)/ (kcatNADP+/KmNADP+) do not change except when trans-2-nonenal is used. This indicates that the interaction with the nicotinamide cofactor is not affected by the aldehyde substrates tested in this analysis (Table 2A). We can conclude that both Thr149 and Val200 are necessary to bind NADP+ efficiently. The Val200 side chain is needed for promoting optimum interactions. It filled the space resulting from positioning the small residue Thr149 instead of Glu on the opposite side. Val200 likely pushed NADP+ in a favorable position to maintain the required contacts with amino acids of the cofactor-binding cleft side-walls. Thr149 added hydrogen bonds to coordinate the 2'-phosphate group. The establishment of such hydrogen bond network surrounding the cofactor not only strengthens with NADP+ but also directs it in a suitable orientation

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which explains the more effective NADP+-related dehydrogenation reaction. This is supported by kinetic studies with a reduced KmNADP+ and a higher catalytic efficiency for ALDH3H1E149T/I200V when compared to other mutated and wild-type enzymes. 3.2.2. Double mutant enzymes carrying a Gly residue at position 200 The Gly residue was introduced because it is present in an identical position in the NADP+-dependent VIBHA_ALDH (Fig. 1). Despite the presence of residues which are capable to coordinate the ribose or the phosphate group at position 149, Km values for both cofactors for all mutated enzymes carrying Gly at position 200 are higher compared to other mutants and the native enzyme (Table 1). Their catalytic efficiencies are very low. This indicates that the overall cofactor-binding ability is affected and neither NAD+- nor NADP+-binding is optimal. Presumably, the absence of a stiff side chain (Val or Ile) affected the widening of the cofactor-binding cleft resulting in a weak stabilization as well as a lack of adjustment of the cofactor inside its binding site, thereby reducing the number of possibilities for interactions. In addition, possible steric clashes can occur with side chains of amino acids of the side-walls of the cofactor-binding pocket which may account for the observed low efficiencies for both cofactors. 3.3. ALDH3H1 with mutations at positions 149, 178 and 200 ALDH3H1E149T/I200V is the most efficient mutant generated in this study when evaluated by improved catalysis with NADP+ (reflected by highest turnover number and catalytic efficiency value). Nonetheless, ALDH3H1E149T/I200V was less effective than naturally occurring homologs preferring NADP+ (human) or NADP+-dependent ALDHs (VIBHA_ALDH) [52]. Therefore, this mutant was selected for further engineering in regions distant from the adenine ribose binding cavity. The comparison of the predicted topology of the cofactor-binding domains of the NAD+-dependent ALDH3H1 and the fatty aldehyde dehydrogenase from V. harveyi (VIBHA_ALDH, solved, PDB: 1EYY) with a high affinity for NADP+ [52] reveals a striking difference (Fig. 5). The molecular surface of the cofactor-binding region of V. harveyi enzyme has a “closed” cleft where Arg210 appears as a key residue in the interaction with the cofactor (Fig. 5). The side chain of Arg210 is positioned over the adenine ring and exerts a stacking force holding the cofactor in an optimal position within the cleft thus facilitating its interaction with other amino acids and resulting in the observed specificity for NADP+ [51]. This is a unique structural feature which is absent in all other NAD(P)+-specific or NAD+-dependent ALDHs. A comparison with the predicted structure of the cofactor-binding site of the NAD+dependent ALDH3H1 reveals an important structural feature that may mediate cofactor-specificity as well as affinity. Arabidopsis ALDH3H1 displays an “open” cleft lacking Arg or any aromatic residue in an

689

identical location exerting a cation-π or π-π stacking interaction on the adenine ring (Fig. 5). The results obtained demonstrate that the introduction of an Arg at this location in the cofactor-binding site (mutation V178R) yielded the enzyme ALDH3H1E149T/V178R/I200V showing an improved affinity for NADP+ (Table 2B). Based on structure homology with the cofactorbinding cleft of VIBHA_ALDH, we suggest that the introduced Arg178 is the structural backbone which improved the binding of NADP+. Its guanidinium group may function as a “gate” exerting a cation-π planar stacking force in the adenine ring of NADP+. It may result in holding the NADP+ buried with an adjusted favorable orientation and pushing the adenosine moiety toward the hydrophobic cavity. It is plausible, that this led to interactions with surrounding amino acids especially by positioning the 2′-phosphate moiety of NADP+ into the adenosyl ribose binding cavity within hydrogen-bonding distance of Thr149 which may explain the low KmNADP+ compared to other mutants. The observed high kcatNADP+ value indicates an improvement in the catalytic mechanism. To confirm the cofactor affinity shift from NAD +-dependency toward NADP+-specificity upon mutating three amino acids simultaneously, dissociation constants for the oxidized nicotinamide cofactors (KdNAD+ and KdNADP+) were determined based on changes of the dehydrogenase activities and cofactor affinities of ALDH3H1 and ALDH3H1E149T/V178R/I200V in the presence or absence of NAD(P)H. The reduced nicotinamide cofactors led to a competitive inhibition when used in the presence of the oxidized forms. This finding is compatible with the kinetic mechanism of most ALDH enzymes. We observed that in the presence of NAD(P)+, KdNADP+ is lower than KdNAD+ for ALDH3H1E149T/V178R/I200V which demonstrates that the binding of NADP+ is favored compared to the binding of NAD+ in the triple mutant. In the presence of NADH, KdNAD+ for ALDH3H1E149T/V178R/I200V is higher than the respective parameter for the wild-type enzyme ALDH3H1. This indicates a weaker binding of NAD+ to the triple mutant enzyme. The increase of concentrations of NADPH resulted in a progressive inactivation of ALDH3H1E149T/V178R/I200V, and KdNAD+ and KdNADP+ are higher than the respective kinetic constants KmNAD+ and KmNADP+. NADH does not affect the triple mutant activity. In contrast, NADH binds to ALDH3H1 resulting inhibition of the enzyme especially at low concentrations of NAD+. Taken together, these observations reveal an enhanced binding of a coenzyme which has a phosphate group on the 2′ position of the ribose ring for the ALDH3H1E149T/V178R/I200V. The introduction of R178 does not affect the binding of NAD+ in the same manner as NADP+ because the adenine ring of NAD+ is not correctly positioned in the same plane due to the shift caused by the absence of the 2′-phosphate group. Because of this shift, the interaction with the arginine guanidinium moiety is not given. The differences in affinity for NADP+ between the triple mutant ALDH3H1E149T/V178R/I200V and the the NADP+-dependent fatty ALDH from V. harveyi, VIBHA_ALDH [43], can be explained by the presence

Fig. 5. Molecular surface representation of cofactor-binding sites of the solved Vibrio hareyi VIBHA_ALDH (panel A. PDB: 1EYY) [51], models of the wild-type A. thaliana ALDH3H1 (panel B) and the triple mutant ALDH3H1E149T/V178R/I200V (panel C). Amino acid residues assumed to be important in VIBHA_ALDH, equivalent residues in ALDH3H1 and residues mutagenized in ALDH3H1E149T/V178R/I200V are depicted. The ligand (NAP-2 in panel A) and the cofactor (NADP+ in panels B and C) is shown in ball-and-stick models. The predicted structure of ALDH3H1 was generated by the web-based server SWISS-MODEL [53] and rendered using PyMol.

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of other amino acids in VIBHA_ALDH which contribute to a tight interaction and correct orientation of the cofactor, thus resulting in the observed higher affinity. 4. Conclusions Homology modeling and extensive mutant analysis permitted to understand cofactor specificity of the A. thaliana NAD+-dependent ALDH3H1 enzyme. Our study provided evidence that amino acids occupying positions 149, 178 and 200 are critical residues determining nicotinamide cofactor specificity (NAD+ and NADP+). Substitution of these residues relaxed the nicotinamide cofactor specificity by rendering the enzyme able to use NADP+ besides NAD+. Dual coenzyme specificity was achieved by combining the two point-mutations E149T/I200V and the synergistic effect of three mutations (E149T/V178R/I200V) was even larger. The cofactor specificity was converted in favor of NADP+, as the triple mutant ALDH3H1E149T/V178R/I200V uses NADP+ with almost 7-fold higher catalytic efficiency than NAD+. The selection of mutants was based on structural information deduced from homology-based modeling and sequence alignment with other NAD(P)+-dependent dehydrogenases as mentioned in the universal protein repository UniProt. This indicates that the predicted structure of ALDH3H1 is a reliable hypothetical working model until a crystal structure of the holoenzyme will be obtained. Blast of the ALDH3H1 sequence against all listed plant family3 ALDHs available in UniProt database, revealed that only two Picea sitchensis (strika spurce) ALDHs (UniProt: B8LML9 and A9NUC8), two Arabidopsis lyrata ALDHs (UniProt: D7KVK7 and D7KNR8) and the A. thaliana ALDH3F1 (UniProt: Q70E96) have an Ile instead of a conserved Val in the position equivalent to 200 in ALDH3H1 (Fig. 6). Thus, it seems that NAD+-dependent dehydrogenases are evolved from enzymes which are able to use both nicotinamide cofactors and

which can have Glu and Val residues at positions equivalent to 149 and 200 in ALDH3H1 respectively. The Arabidopsis chloroplastic isoform ALDH3I1 [34] may represent the ancestor of the cytosolic isoform ALDH3H1 because it has an Ile instead of Val200 (ALDH3H1 numbering). The NAD+-dependency of ALDH3H1 may have occurred as an adaptation to the environment or the functions required in the cytoplasm. In members of the divergent group of bacterial NADP+-dependent fatty ALDHs (universal protein resource UniProt), position 200 (ALDH3H1 numbering) is occupied by a Gly residue. The Glu residue occupying position 149 in ALDH3H1 is substituted by other residues including Gln, Asp, Asn, Thr, Ser or Pro (Figs. 6 and A.3). The absence of Arg present in V. harveyi ALDH at position equivalent to 178 (ALDH3H1 numbering) seem to be a principal factor which determines the shift from NADP+-dependency toward relaxed nicotinamide cofactor specificity. 5. Materials and methods 5.1. Site-directed mutagenesis Site-directed mutagenesis was performed based on the method of the QuickChange® site-directed mutagenesis kit (Stratagene, La Jolla, CA, USA) using primers carrying the desired mutations, Pfu Turbo® DNA polymerase (Agilent Technologies, Santa Clara, CA, USA) DpnI (Thermo Fisher Scientific, St. Leon-Rot, Germany) [34]. All mutated ALDH3H1 constructs, the corresponding mutagenic primers and positions of substituted amino acid in the recombinant enzymes are listed in Table A.1. Mutants are designated according to the substituted amino acid position in the native protein. First, the recombinant plasmid pET28a-ALDH3H1 [34], was used as template to substitute Ile200 by Val or Gly yielding ALDH3H1I200V or ALDH3H1I200G. In a second round of mutations, the recombinant

Fig. 6. Alignment of amino acid sequences including cofactor-binding regions among different class 3 and NADP+-dependent bacterial fatty aldehyde dehydrogenases. Amino acid sequences correspond to the following plant and bacterial aldehyde dehydrogenase enzymes: Arabidopsis thaliana ALDH3H1 (UniProt: Q70DU8), ALDH3F1 (UniProt: Q70E96) and ALDH3I1 (UniProt: Q8W033), A. lyrata subsp. Lyrata ARALL_AL1 (UniProt: D7KNR8) and ARALL_AL2 (UniProt: D7KVK7), Picea sitchensis PICSI_AL1 (UniProt: B8LML9) and PICSI_AL2 (UniProt: A9NUC8), Medicago truncatula MEDTR_ALDH (UniProt: B7FM69), Populus trichocarpa POPTR_AL1 (UniProt: B9H7G3) and POPTR_AL2 (UniProt: B9GU23), Ricinus communis RICCO_ALDH (UniProt: B9S2Y3), Glycine max SOYBN_ALDH (UniProt: I1JZ49), Vitis vinifera VITVI_ALDH (UniProt: D7SWG5), Craterostigma plantagineum Cp-ALDH (UniProt: Q8VXQ2), Zea mays MAIZE_ALDH (UniProt: B4FUA7), Oryza sativa subsp. japonica ORYSJ_ALDH (UniProt: Q53NG8), Oryza sativa subsp. Indica ORYSI_ALDH (UniProt: B8BJI2), bacterial NADP+dependent fatty aldehyde dehydrogenases as mentioned in UniProt databse: Mycobacterium smegmatis MYCS2_ALDH (UniProt: A0R5B7), Corynebacterium ammoniagenes CORAM _ALDH (UniProt: D5P1C2), Ralstonia metallidurans RALME_ALDH (UniProt: Q1LQ58), Silicibacter sp. TrichCH4B; SILIC_ALDH (UniProt: C9D1A0), Vibrio harveyi VIBHA _ALDH (UniProt: Q56694). Alignment was performed using ClustalW web-based multiple sequence alignment tool. Conserved residues are shaded in black. Residues of cofactor-binding site which were targeted for site-directed mutagenesis are indicated by signs (♣, ♠ and ♦). Residues occupying these positions are highlighted with different colors. Conserved motifs involved in the catalytic process (motifs 1 and 5) and the cofactor binding (motifs 2 and 4) are boxed. Numbers in blue on the left-hand and right-hand side for each sequence indicate the positions of the amino acids in each native protein. Amino acids underlined in red indicate the characteristic glycine rich sequence. The 2D structure diagram on the top corresponds to a prediction deduced from the structure homology based model of Arabidopsis ALDH3H1 (performed using SWISS-MODEL using ALDH3a1, PDB: 3SZB, as template), where α-helices and β-strands are labeled as stated in the convention used for alcohol dehydrogenases.

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plasmid pET28a-ALDH3H1 and resulting plasmids from the first mutagenic experiment were used as templates to substitute Glu149 by Gln, Asn, Asp or Thr and combine mutations at positions 149 and 200 respectively. Amino acid residues Gln, Asn, Asp and Thr were chosen to substitute Glu149 because they are present in an equivalent position in bacterial NADP+-dependent fatty aldehyde dehydrogenases as cited in the universal protein database UniProt (Fig. 6). In these residues the length of the side chain or/and the charge are different from the initial Glu residue in position 149. The resulting mutated plasmid (pET28a-ALDH3H1E149T/I200V) carrying two mutations was used to combine all mutations at amino acid positions 149, 178 and 200 after introducing Arg instead of Val178. All mutated plasmids were transformed into Escherichia coli DH10B (Invitrogen, Carlsbad, CA, USA) and verified by DNA sequencing and finally transformed into E. coli BL21 (DE3) (New England Biolabs Inc., Ipswich, MA, USA) to express active enzymes.

5.2. Expression and purification of recombinant enzymes The pET28a-ALDH3H1 plasmid [34] and the mutated derivatives were used for the overexpression of the N-terminal hexahistidinetagged recombinant enzymes. Cultures of E. coli BL21 (DE3) cells transformed with the plasmids were grown in LB media supplemented with 50 μg/mL kanamycin at 37 °C. When the log phase was reached (OD600 ≈ 0.500), expression of recombinant proteins was induced by adding IPTG (isopropyl β-D-thiogalactopyranoside) to a final concentration of 0.1 mM. Cultures were further incubated at 24 °C with optimal aeration (shaking at 250 rpm) for 3 h. Cells were harvested by centrifugation at 5300 rpm, 4 °C, for 20 min and stored at −20 °C until use. Purification of recombinant proteins was performed as described [34]. Cell pellets were thawed on ice and resuspended in 5 mL lysis buffer (50 mM Hepes/NaOH (pH 7.4), 300 mM NaCl, 10% (v/v) glycerol, 0.1% (v/v) Triton X-100, 1.5 mM β-mercaptoethanol; freshly added, 5 mM imidazole, 1 mg/mL lysozyme). The frozen cell suspension was further lyzed by sonication (6 bursts of 30 s followed each by intervals of 2 min for cooling) using ultrasonic disintegrator UP200S (Hielscher Ulrasonics GmbH, Teltow, Germany). The cell lysate was centrifuged at 14000 rpm at 4 °C for 30 min, and the supernatant containing the crude extract was filtered through a 0.45 μm filter and used for affinity chromatography. For affinity chromatography, columns packed with HIS-Select® Nickel Affinity Gel were used (Sigma-Aldrich, St. Louis, MO, USA) and equilibrated with 3 times the column volume of lysis buffer without lysozyme (see above). The clarified filtrate was passed two times through the resin. Columns were washed with 10 times the column volume of the lysis buffer without lysozyme followed by a washing step with lysis buffer without lysoszyme containing 20 mM imidazole before eluting the His-tagged enzyme in 12 fractions of 250 μL using lysis buffer without lysozyme containing 250 mM imidazole. Fractions corresponding to the elution peak were selected and adjusted to 50% (v/v) glycerol, 1 mM PMSF, 0.5 mM NAD+ and 10 mM DTT to stabilize the enzymes before storage at −80 °C. Corresponding NAD+-related ALDH activities of all eluted fractions were assayed using 5 μL of each eluted fraction (Fig. A.1).

5.3. Recombinant enzyme characterization Protein concentrations were determined using the Bradford assay (Bio-Rad, Munich, Germany) with bovine serum albumin as reference protein. The purity of the enzymes was verified by analyzing them in non-reducing 10% SDS/PAGE. Gels were stained with Colloidal Coomassie blue (Thermo Fisher Scientific, St. Leon-Rot, Germany) to visualize the proteins (Fig. A.1). Corresponding NAD+-related ALDH activities of all eluted fractions were also assayed.

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The web-based tool ExPASy compute pI/MW (Swiss Institute of Bioinformatics, Geneva, Switzerland; http://web.expasy.org/compute_pi/) was used to estimate the molecular weight of the proteins. SDS/PAGE electrophoresis shows that all the purified ALDHs form a prominent band of about 56 kDa which corresponds to the predicted molecular weight. Gel image analysis was performed using imageJ software version 1.46 (National Institute of Health, Bethesda, MD, USA), it showed that only minor protein contaminants (b5%) can be detected in some eluted fractions. 5.4. Homology modeling The A. thaliana dehydrogenase ALDH3H1 model was built using the web-based structure homology-modeling server SWISS-MODEL [54]. The solved three dimensional structures of rat ALDH3a1 (UniProt: P11883, PDB: 1AD3) [14] and human ALDH3A1 (UniProt: P30838, PDB: 3SZB) [15] were used as templates; they exhibit respectively 45% and 46% identity with the recombinant Arabidopsis ALDH3H1 respectively. The predicted structures obtained were rendered using the molecular visualization tool PyMol as a PDB viewer (DeLano Scientific; http://pymol.org) and then visually inspected for obvious inaccuracies. The reliability of the generated models using the SWISS-MODEL tool was checked and they were rebuilt using other comparative modeling methods such as I-tasser [54]. Predicted structures were compared and the topologies of the generated cofactor binding sites were identical. The Pymol tool was also used to simulate the performed mutations as well as estimating distances between amino acid side chains across the initial and mutated cofactor-binding clefts. AutoDock/Vina program [55] was used to simulate the binding of NAD(P)+ to the enzyme. Sequence alignments were performed using ClustalW2 (EMBL, European Bioinformatics Institute) [56]. 5.5. Enzyme assays and determination of enzymatic parameters Activities of ALDH enzymes were determined spectrophotometrically by measuring the absorbance at λ = 340 nm reflecting the amount of NAD(P)H produced upon reduction of NAD(P)+ (εNAD(P)H = 6.22 mM−1.cm−1 at λ = 340 nm) [34]. All enzymatic assays were performed at 25 °C using a GENESYS 10 UV spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). The reactions were started by adding 1 μg of the enzyme protein to the assay buffer containing 100 mM sodium pyrophosphate at pH 8 which corresponds to the pH optimum for ALDH3H1 enzymatic activity [34] supplemented with NAD+ (Roche Applied Science, Mannheim, Germany) or NADP+ (Sigma-Aldrich, St. Louis, MO, USA) as cofactor and aliphatic aldehydes as substrate at the corresponding saturating concentrations for each ALDH3H1 mutated isoform. Substrate saturating concentrations were determined by a series of assays where values of enzymatic activities were plotted versus varied concentrations of substrate in the presence of NAD(P)+ at a fixed concentration corresponding to expected enzyme saturation. For the assessment of kinetic parameters relative to cofactor specificity, mutated enzymes were analyzed by a matrix of assays in which both cofactor concentrations were varied over a broad range (12.5 μM to 10 mM) in the presence of substrates at saturating concentrations (Fig. A.2). The data resulted in a Michaelis–Menten saturation curve where activities were determined as functions of either cofactor concentration. Hexanal (Sigma-Aldrich, St. Louis, MO, USA) was used as substrate in all initial assays because of its higher solubility in aqueous solutions. Other aliphatic aldehyde substrates, i.e., trans-2-hexenal, nonanal, trans-2-nonenal (Sigma-Aldrich, St. Louis, MO, USA) were used in addition as indicated. Enzyme specificities for cofactors and substrates are reported as kcatNAD(P)+/KmNAD(P)+ (s−1.mM−1). Related specific activities are initially calculated as Vmax (μmol NAD(P)H.min−1.mg−1) for

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the used enzyme, then converted into the turnover number kcat (s−1). Catalytic efficiencies are reported as kcat/Km (app)(s−1.mM−1). All assays were performed at least in independent triplicates. All kinetic parameters are reported as mean values ± S.E.M. Equilibrium dissociation constants for both cofactors (KdNAD+ and KdNADP+) were determined by following changes of the reaction rate as well as affinities of cofactors upon addition of NAD(P)H. Enzymatic reactions were carried out using hexanal at saturated concentrations. Concentrations of NADH or NADPH were kept constant for each set of experiments. The concentration of NAD(P)H was different within different experimental series (from 10 μM to 375 μM) depending on the assayed enzyme and the cofactor. Data were analyzed using nonlinear regression of the Michaelis–Menten equation to determine the equilibrium dissociation constants. Calculations were performed from at least three independent series of measurements and mean values ± S.E.M were used for subsequent interpretations. In the Michaelis Menten equation, V = V max [NAD(P) + ]/K mNAD(P) + + [NAD(P) + ] where KmNAD(P)+ = (k2 + k9)/k1 and KdNAD(P)+ = k2/k1. Thus, KmNAD(P)+ =KdNAD(P)+ + k9/k1 (Fig. A.4). An abortive binary complex EnzymeNAD(P)H may form upon addition of NAD(P)H, consequently, the catalysis will slow down and k 9 will be negligible. Accordingly, we can consider that KmNAD(P)+ corresponds to KdNAD(P)+. Therefore the equilibrium dissociation constant values were determined from the Michaelis–Menten plots which represent the concentration of NAD(P) + corresponding to half of the maximum rate of the reaction. Supplementary data to this article can be found online at http://dx. doi.org/10.1016/j.bbapap.2014.01.008. Acknowledgements We gratefully acknowledge Valentino Giarola, IMBIO, University of Bonn for helping with the phylogenetic analysis. References [1] J. Perozich, H. Nicholas, B.C. Wang, R. Lindahl, J. Hempel, Relationships within the aldehyde dehydrogenase extended family, Protein Sci. 8 (1999) 137–146. [2] H.H. Kirch, D. Bartels, Y. Wie, P.S. Schnable, A.J. Wood, The ALDH superfamily of Arabidopsis, Nucleic Acids Res. 9 (2004) 371–377. [3] C. Brocker, M. Vasiliou, S. Carpenter, C. Carpenter, Y. Zhang, X. Wang, S.O. Kotchoni, A.J. Wood, H.H. Kirch, D. Kopečný, D.W. Nebert, V. Vasiliou, Aldehyde dehydrogenase (ALDH) superfamily in plants: gene nomenclature and comparative genomics, Planta 237 (2012) 189–210. [4] N.A. Sophos, V. Vasiliou, Aldehyde dehydrogenase gene superfamily: the 2002 update, Chem. Biol. Interact. 143–144 (2003) 5–22. [5] B. Jackson, C. Brocker, D.C. Thompson, W. Black, K. Vasiliou, D.W. Nebert, V. Vasiliou, Update on the aldehyde dehydrogenase gene (ALDH) superfamily, Hum. Genomics 5 (2011) 283–303. [6] C. Gao, B. Han, Evolutionary and expression study of the aldehyde dehydrogenase (ALDH) gene superfamily in rice (Oryza sativa), Gene 431 (2009) 86–94. [7] Y. Zhang, L. Mao, H. Wang, C. Brocker, X. Yin, V. Vasiliou, Z. Fei, X. Wang, Genome-wide identification and analysis of grape aldehyde dehydrogenase (ALDH) gene superfamily, PLoS One 7 (2012) e32153. [8] S.O. Kotchoni, J.C. Jimenez-Lopez, A.P. Kayodé, E.W. Gachomo, L. Baba-Moussa, The soybean aldehyde dehydrogenase (ALDH) protein superfamily, Gene 495 (2012) 128–133. [9] A.J. Wood, R.J. Duff, The aldehyde dehydrogenase (ALDH) gene superfamily of the moss Physcomitrella patens and the algae Chlamydomonas reinhardtii and Ostreococcus tauri, Bryologist 112 (2009) 1–11. [10] S.A. Moore, H.M. Baker, T.J. Blythe, K.E. Kitson, T.M. Kitson, E.N. Baker, Sheep liver cytosolic ALDH: the structure reveals the basis for retinal specificity, Structure 6 (1998) 1541–1551. [11] C.G. Steinmetz, P. Xie, H. Weiner, T.D. Hurley, Structure of mitochondrial ALDH: the genetic component of ethanol aversion, Structure 5 (1997) 701–711. [12] W.K. Tang, K.B. Wong, Y.M. Lam, S.S. Cha, C.H. Cheng, W.P. Fong, The crystal structure of seabream antiquitin reveals the structural basis of its substrate specificity, FEBS Lett. 582 (2008) 3090–3096. [13] A.G. Díaz-Sánchez, L. González-Segura, E. Rudiño-Piñera, A. Lira-Rocha, A. Torres-Larios, R.A. Muñoz-Clares, Novel NADPH-cysteine covalent adduct found in the active site of an aldehyde dehydrogenase, Biochem. J. 439 (2011) 443–452. [14] Z.J. Liu, Y.J. Sun, J. Rose, Y.J. Chung, C.D. Hsiao, W.R. Chang, I. Kuo, J. Perozich, R. Lindahl, J. Hempel, B.C. Wang, The first structure of an aldehyde dehydrogenase reveals novel interactions between NAD and the Rossmann fold, Nat. Struct. Biol. 4 (1997) 317–326.

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