Inhibition, crystal structures, and in-solution oligomeric structure of aldehyde dehydrogenase 9A1

Article abstract of DOI:10.1016/j.abb.2020.108477

Aldehyde dehydrogenase 9A1 (ALDH9A1) is a human enzyme that catalyzes the NAD⁺-dependent oxidation of the carnitine precursor 4-trimethylaminobutyraldehyde to 4-N-trimethylaminobutyrate. Here we show that the broad-spectrum ALDH inhibitor diethylaminobenzaldehyde (DEAB) reversibly inhibits ALDH9A1 in a time-dependent manner. Possible mechanisms of inhibition include covalent reversible inactivation involving the thiohemiacetal intermediate and slow, tight-binding inhibition. Two crystal structures of ALDH9A1 are reported, including the first of the enzyme complexed with NAD⁺. One of the structures reveals the active conformation of the enzyme, in which the Rossmann dinucleotide-binding domain is fully ordered and the inter-domain linker adopts the canonical β-hairpin observed in other ALDH structures. The oligomeric structure of ALDH9A1 was investigated using analytical ultracentrifugation, small-angle X-ray scattering, and negative stain electron microscopy. These data show that ALDH9A1 forms the classic ALDH superfamily dimer-of-dimers tetramer in solution. Our results suggest that the presence of an aldehyde substrate and NAD⁺ promotes isomerization of the enzyme into the active conformation.

Full text of DOI:10.1016/j.abb.2020.108477

Journal Pre-proof
Inhibition, crystal structures, and in-solution oligomeric structure of aldehyde
dehydrogenase 9A1
Jesse W. Wyatt, David A. Korasick, Insaf A. Qureshi, Ashley C. Campbell, Kent S.
Gates, John J. Tanner
PII:
S0003-9861(20)30486-0
https://doi.org/10.1016/j.abb.2020.108477
YABBI 108477
DOI:
Reference:
To appear in: Archives of Biochemistry and Biophysics
Received Date: 23 May 2020
Revised Date: 19 June 2020
Accepted Date: 20 June 2020
Please cite this article as: J.W. Wyatt, D.A. Korasick, I.A. Qureshi, A.C. Campbell, K.S. Gates, J.J.
Tanner, Inhibition, crystal structures, and in-solution oligomeric structure of aldehyde dehydrogenase
9A1, Archives of Biochemistry and Biophysics (2020), doi: https://doi.org/10.1016/j.abb.2020.108477.
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Inhibition, crystal structures, and in-solution oligomeric structure of aldehyde
dehydrogenase 9A1
a
b
c
b
a,b
Jesse W. Wyatt , David A. Korasick , Insaf A. Qureshi, Ashley C. Campbell, Kent S. Gates ,
b,a,*
and John J. Tanner
a
Department of Chemistry, University of Missouri, Columbia, Missouri 65211, United States
b
Department of Biochemistry, University of Missouri, Columbia, Missouri 65211, United States
c
Department of Biotechnology & Bioinformatics, School of Life Sciences, University of
Hyderabad, Prof. C.R. Rao Road, Hyderabad, 500046, India.
*
Corresponding author: Department of Biochemistry, University of Missouri, Columbia, MO
6
5211. Phone: (573) 884-1280. E-mail: tannerjj@missouri.edu.
Running Title: Structure of ALDH9A1
Databases: Coordinates and structure factor amplitudes have been deposited in the Protein Data
Bank under accession codes 6VR6 and 6VWF. SAXS data sets have been deposited in the
Small-Angle Scattering Biological Data Bank under the following accession codes: SASDHV5,
SASDHW5, and SASDHX5.
Conflict of Interest: The authors declare no competing financial interest.
1

Abstract
+
Aldehyde dehydrogenase 9A1 (ALDH9A1) is a human enzyme that catalyzes the NAD -
dependent oxidation of the carnitine precursor 4-trimethylaminobutyraldehyde to 4-N-
trimethylaminobutyrate. Here we show that the broad-spectrum ALDH inhibitor
diethylaminobenzaldehyde (DEAB) reversibly inhibits ALDH9A1 in a time-dependent manner.
Possible mechanisms of inhibition include covalent reversible inactivation involving the
thiohemiacetal intermediate and slow, tight-binding inhibition. Two crystal structures of
+
ALDH9A1 are reported, including the first of the enzyme complexed with NAD . One of the
structures reveals the active conformation of the enzyme, in which the Rossmann dinucleotide-
binding domain is fully ordered and the inter-domain linker adopts the canonical β-hairpin
observed in other ALDH structures. The oligomeric structure of ALDH9A1 was investigated
using analytical ultracentrifugation, small-angle X-ray scattering, and negative stain electron
microscopy. These data show that ALDH9A1 forms the classic ALDH superfamily dimer-of-
dimers tetramer in solution. Our results suggest that the presence of an aldehyde substrate and
+
NAD promotes isomerization of the enzyme into the active conformation.
Keywords: aldehyde dehydrogenase; ALDH9A1; X-ray crystallography; small-angle X-ray
scattering; analytical ultracentrifugation; negative-stain electron microscopy; time-dependent
inhibition; covalent reversible inhibitor; diethylaminobenzaldehyde; DEAB.
Abbreviations: ALDH, aldehyde dehydrogenase; ALDH9A1, aldehyde dehydrogenase 9A1;
DEAB, diethylaminobenzaldehyde; IMAC, immobilized metal-affinity chromatography; SAXS,
small-angle X-ray scattering; SEC, size-exclusion chromatography; TCEP, tris(2-
carboxyethyl)phosphine; TMBAL, 4-trimethylaminobutyraldehyde.
2

1
. Introduction
The aldehyde dehydrogenase (ALDH) structural superfamily is a large group of enzymes that
+
catalyze the NAD -dependent oxidation of aldehydes to carboxylic acids [1, 2]. The superfamily
comprises hundreds of distinct genes, including 19 ALDHs expressed in humans. ALDHs share
a common protein fold and catalytic mechanism, but subtle differences in their active sites result
in different preferences for the aldehyde substrate.
+
ALDH9A1 is a human enzyme that catalyzes the NAD -dependent oxidation of a variety of
aldehydes including the carnitine precursor 4-trimethylaminobutyraldehyde (TMBAL, Scheme
1
) [3], the GABA precursor aminobutyraldehyde [4], the dopamine metabolite 3,4-
dihydroxyphenylacetaldehyde [5], and betaine aldehyde [6]. A recent comprehensive study of
substrate specificity showed that ALDH9A1 has a strong (> 10-fold) preference for TMBAL [7],
confirming the assertion of Vaz, et al. [3] that the major in vivo function of this enzyme is to
catalyze the penultimate step of carnitine biosynthesis (E.C. 1.2.1.47), the oxidation of TMBAL
to 4-N-trimethylaminobutyrate. Carnitine functions in the transport of long-chain fatty acids
from the cytosol to the mitochondrial matrix for the synthesis of acyl-CoAs for β-oxidation.
Because the mitochondrial membrane is impermeable to acyl-CoAs, fatty acids are conjugated to
carnitine to enter mitochondria. Thus, ALDH9A1 functions indirectly in β-oxidation.
Recently the first crystal structure of human ALDH9A1 was reported [7] (PDB ID: 6QAP).
Although ALDH9A1 exhibits the basic ALDH superfamily fold, the structure revealed two
remarkable features. First, the final α-helix and β-strand of the Rossmann dinucleotide-binding
fold are disordered. Referred to as αE-βE in the closely-related betaine ALDH [8], these
+
secondary structural elements form extensive interactions with NAD in other ALDHs. Further,
αE forms a major part of the dimer interface in other ALDHs. The second atypical aspect of the
3

apo ALDH9A1 structure concerns the ~25-residue inter-domain linker, which connects the
catalytic domain to the oligomerization domain. In other ALDH structures, the linker folds into a
long β-hairpin, wherein the strands of the β-hairpin form the floor of the aldehyde substrate
+
tunnel, while the tip helps stabilize the NAD binding site. In contrast, the linker in the apo
ALDH9A1 structure adopts a novel S-shaped conformation, which blocks access to the aldehyde
substrate binding pocket. The apo ALDH9A1 structures appear to represent an inactive enzyme,
raising the question of how αE-βE and the inter-domain linker isomerize into their active
conformations.
Herein we describe a structural and biochemical study of ALDH9A1 designed to provide
understanding of the catalytic function of this enzyme, in light of the atypical structural features
observed previously for the apo enzyme. We demonstrate that diethylaminobenzaldehyde
(DEAB) is a reversible, time-dependent inhibitor of ALDH9A1. We report two crystal structures
+
of ALDH9A1 complexed with NAD . In one of the structures, which was obtained from enzyme
+
treated with DEAB and NAD , αE-βE and the inter-domain linker are ordered and adopt the
canonical secondary, tertiary, and quaternary structures observed in other ALDHs. In contrast,
+
the other NAD complex structure exhibits the atypical features of the apo enzyme, showing that
+
NAD binding alone is not sufficient to promote the active conformation of the enzyme. The
oligomeric structure of ALDH9A1 was determined using analytical ultracentrifugation, small-
angle X-ray scattering, and negative stain electron microscopy. These studies show that
ALDH9A1 forms the classic ALDH superfamily dimer-of-dimers tetramer in solution. Our
results provide insight into the mechanism by which ALDH9A1 is activated for catalysis.
4

2
. Materials and methods
2
.1. Expression plasmids
A synthetic gene encoding human ALDH9A1 with codons optimized for expression in
Escherichia coli was purchased from Genscript. The gene was subcloned by Genscript into
plasmid pET-24b(+) between NdeI/XhoI restriction sites. The expressed protein contains an N-
terminal hexahistidine tag and tobacco etch virus protease cleavage site.
Removal of the His-tag from the protein expressed from the aforementioned pET-24b(+)
construct was problematic, so another expression plasmid encoding ALDH9A1 fused to SUMO
was created. The coding sequence of human ALDH9A1 was PCR amplified from the pET24b(+)
construct, gel-purified, and cloned into pET-SUMO using BamHI and XhoI restriction sites.
Positive clones were identified by colony PCR followed by DNA sequencing using T7 forward
and reverse primers.
2
.2. Purification of SUMO-ALDH9A1 and His-ALDH9A1
To purify SUMO-tagged ALDH9A1, the fusion protein was over-expressed in E. coli BL21
(DE3) with induction by 0.25 mM IPTG at 18°C for 20 hrs. Cells were collected by
centrifugation and resuspended in buffer A (50 mM Tris pH 8.0, 300 mM NaCl, 10 mM
imidazole and 5% glycerol (v/v) containing 0.1% Triton X-100. Cells were lysed by sonication
and then centrifuged at 20,000 rpm in a SS-34 rotor for 30 min at 4ºC to remove the remaining
insoluble material. The supernatant was subjected to immobilized metal-affinity chromatography
(
IMAC) on a HisTrap HP column (GE Healthcare) charged with nickel ion and equilibrated with
buffer A and unbound proteins were washed sequentially with ten column volumes each of
buffer A with imidazole concentrations of 40, 100 and 150 mM. The His-tagged proteins were
5

eluted with buffer A containing 200 mM imidazole. To remove the N-terminal SUMO-tag, His-
tagged SUMO protease (purified separately by IMAC chromatography) was added to IMAC-
purified His-SUMO-ALDH9A1 in a 1:100 ratio and subjected to dialysis for 12 – 16 h at 4°C in
2
5 mM Tris-HCl pH 8.0, 100 mM NaCl, 5% (v/v) glycerol and 0.5% Tris(2-
carboxyethyl)phosphine (TCEP). The digested protein sample was then passed through the
IMAC column, which retained the cleaved SUMO tag, SUMO protease, and uncleaved fusion
protein. The pure, untagged ALDH9A1 protein was collected in the flow through and then
protein quality was analyzed with SDS-PAGE. Subsequently, ALDH9A1 was concentrated using
a 50 kDa MilliporeSigma Amicon Ultra centrifugal concentrator and purified further using size-
exclusion chromatography (SEC) on a HiLoad 16/60 Superdex 200 column with a buffer
containing 50 mM Tris-HCl pH 8.0, 600 mM NaCl, 5% (v/v) glycerol and 0.5% TCEP. The
purity of the recombinant ALDH9A1 was verified by SDS-PAGE. Protein concentration was
determined by absorbance using an estimated extinction coefficient calculated by ProtParam [9].
His-tagged ALDH9A1 was over-expressed from the pET-24b(+) vector in E. coli BL21
(DE3) at OD ~ 0.6 with 0.25 mM IPTG at 18°C for 18 – 21 hrs. Cells were harvested by
centrifugation and resuspended in buffer A (50 mM HEPES pH 8.0, 300 mM NaCl, 10 mM
imidazole, and 5% glycerol) containing 0.1% Triton X-100 and 0.4 mM PMSF. Cells were lysed
by sonication and centrifuged at 16,500 rpm in a SS-34 rotor for 1 h at 4°C. The supernatant was
subjected to IMAC using a HisTrap HP column (GE Healthcare) charged with nickel ion and
equilibrated in buffer A. After washing the column with 100 mM imidazole, His-ALDH9A1 was
eluted with 200 mM imidazole. The enzyme was concentrated with a 50 kDa MilliporeSigma
Amicon Ultra centrifugal concentrator and further purified by SEC on a Superdex 200 column in
the presence of 50 mM HEPES pH 8.0, 100 mM NaCl, 5% (v/v) glycerol, and 0.5 mM
6

dithiothreitol (DTT). The purity of His-ALDH9A1 was verified with SDS-PAGE and protein
concentration was determined by A280 using a NanoDrop 2000c (ThermoFisher) and confirmed
via bicinchoninic acid assay (Pierce).
2
.3. Kinetics of enzyme activity and inhibition
All kinetic measurements were performed using His-tag-free ALDH9A1. The steady state
+
kinetic parameters of ALDH9A1 for the aldehyde substrate hexanal at saturating NAD
concentration were determined. These assays monitored NADH production (340 nm) at 27°C in
an Epoch 2 plate reader (BioTek, Winooski, VT, USA). The assay buffer contained 100 mM
sodium pyrophosphate pH 8.0. The aldehyde substrate concentration was varied in the range of
+
2
.5 – 150 µM, while the NAD concentration was fixed at 1.5 mM. The untagged enzyme was
used at 500 nM. Each measurement was performed in triplicate. Initial rates were estimated by
linear regression of the first five minutes of the assay. The absorbance at 340 nm was converted
-1
-1
to NADH concentration using an extinction coefficient of 6220 M cm . Kinetic parameters
were obtained by fitting the initial rate data to the Michaelis-Menten equation using Origin
version 2019 (Fig. S1 of Supplementary data).
DEAB is a substrate for some ALDHs, so we first tested this possibility for ALDH9A1 using
the strategy of Hurley’s group [10]. In this experiment, ALDH9A1 is combined with DEAB and
+
NAD , and then the absorbance at 360 nm and 300 nm are monitored. A decrease at 360 nm
indicates the consumption of DEAB, while an increase at 300 nm indicates the production of the
carboxylic acid product, diethylaminobenzoic acid. For these experiments, the final assay
+
mixture (total volume of 1 mL) contained 0.5 µM ALDH9A1, 30 µM DEAB, and 1.5 mM NAD
in a buffer containing 100 mM sodium pyrophosphate pH 8.0 and 0.1% DMSO. The reaction
7

was initiated by the addition of enzyme. The absorbance was measured for 240 minutes using the
cuvette port of a NanoDrop 2000c (ThermoFisher). Scans covering the wavelength range of 270
–
460 nm were recorded every 10 minutes.
The time-dependent inhibition of ALDH9A1 by DEAB was studied using a progress curve
approach [11, 12]. Progress curve experiments were performed in a BioTek Epoch 2 plate reader
in duplicate at 27°C with an assay mixture consisting of 0.5 µM ALDH9A1, 2 mM hexanal, 2.5
+
mM NAD , 2 – 5.0 µM DEAB, 0.04% (v/v) DMSO, and 100 mM sodium pyrophosphate pH
8
.0. The total reaction assay volume was 200 µL. The reaction was initiated by the addition of an
+
aliquot of an enzyme- NAD solution and then monitored by observing the production of NADH
at 340 nm. The progress curves measured at different DEAB concentrations were analyzed using
global fitting to combined Equations 1a and 1b [13].
ꢉꢌ − ꢉꢊ
ꢎ1 − ꢈꢏꢐꢑꢒ + ꢅꢓ (1ꢅ)
ꢀ
ꢁꢂꢃꢄꢁꢅꢆꢇꢈ = ꢉ ꢋ +
ꢊ
ꢍ
ꢎꢖꢒ/ꢗꢘ
ꢍ = ꢍ + ꢍ
(1ꢁ)
ꢔ
ꢕ
ꢎꢚꢒ ꢎꢖꢒ
ꢙ1 +
+
ꢜ
ꢗꢛ
ꢗꢘ
Equations 1a and 1b describe the inhibition model in Scheme 2B (see equations 6 and 9 of
Morrison and Walsh [13]). The parameters v and v are the initial and steady-state velocities, t is
i
s
time, a is the initial absorbance offset parameter, and k the apparent first-order rate constant for
o
the establishment of the equilibrium between E·I and E·I*. In Equation 1b, k and k are the
5
6
*
forward and reverse rate constants for the isomerization of E·I to E·I , respectively; K is
I
dissociation constant for the E·I complex; [I] is the inhibitor concentration; [S] is the hexanal
substrate concentration; and K is the Michaelis constant for hexanal, which was estimated to be
m
6
.3 µM in a separate experiment performed in the absence of DEAB (Fig. S1 of Supplementary
8

5 6 I
data). Global fitting was done with Origin 2019, with k , k , and K considered to be global
parameters (shared by all data sets), while K , [S], and [I] were fixed at their known values.
m
-5
-1
-1
During fitting, k refined to a very small value (2 x 10 min ) compared to k (0.2 min );
6
5
however, the uncertainty of k was large, which prevented convergence. Therefore, k was fixed
6
6
-5
-1
at 2 x 10 min to aid convergence of the fitting algorithm.
+
2
.4. Crystallization of ALDH9A1-NAD in space group P1
+
-
Prior to crystallization, DEAB and NAD were added to the protein stock solution (6 mg mL
1
tag-free ALDH9A1; 50 mM Tris-HCL pH 8.0, 600 mM NaCl, 5% glycerol, and 0.5% TCEP)
to final concentrations of 5 mM and 10 mM, respectively. Initial crystal screening was performed
in 96-well Swissci 2 drop MRC Crystallization plates with Hampton Index at 8°C. Promising
crystals were obtained in a condition containing 0.2 M NaCl, 0.1 M Bis-Tris pH 6.5, and 25%
(w/v) PEG 3350. Further improvement was obtained using an additive screen approach. In this
approach, the reservoir consisted of equal volumes of the aforementioned base condition (0.2 M
NaCl, 0.1 M Bis-Tris pH 6.5, and 25% (w/v) PEG 3350) and an additional crystal screen
condition. Improvement in the crystal quality was observed when using Hampton Index
condition G8 as the additive (0.2 M ammonium acetate, 0.1 M HEPES pH 9.5, and 25% (w/v)
PEG 3350). Thus, the final reservoir solution contained 0.1 M NaCl, 0.05 M Bis-Tris 6.5, 0.1 M
ammonium acetate, 0.05 M HEPES pH 9.5, and 25% (w/v) PEG 3350. The crystals were
cryoprotected with 0.1 M NaCl, 0.05 M Bis-Tris pH 6.5, 0.1 M ammonium acetate, 0.05 M
HEPES pH 9.5, 25% (w/v) PEG 3350, 20% (v/v) hexylene glycol, 2.5 mM DEAB and 5 mM
+
NAD . Although the presence of DEAB in the active site was not obvious from the electron
density, its inclusion in the crystallization protocol apparently facilitated growth of a new crystal
9

form, which revealed the active conformation of the enzyme.
+
2
.5. Crystallization of ALDH9A1-NAD in space group C222
+
-1
Prior to crystallization, NAD was added to the protein stock solution (10 mg mL His-
tagged ALDH9A1; 50 mM HEPES pH 8.0, 100 mM NaCl, 5% (v/v) glycerol, and 0.5 mM DTT)
to a final concentration of 30 mM. A crystal screen was performed in 96-well Swissci 2 drop
MRC Crystallization plates at 8°C using Hampton Index with a drop ratio of 0.6/0.4
(enzyme/reservoir). Microseed matrix screening [14, 15] was used with a seed stock prepared
from previously grown C222 crystals. The crystal used for data collection was harvested directly
from Hampton Index condition H5 (0.1 M succinic acid pH 7.0, 15% (w/v) PEG 3350) without
optimization and cryoprotected with the reservoir supplemented with 20% (v/v) hexylene glycol.
2
.6. Crystal structure determination
X-ray diffraction data from a P1 crystal were collected at NECAT beamline ID-E of the
Advanced Photon Source using an EIGER 16M detector in shutterless mode. The dataset used
for refinement consisted of 700 frames spanning a rotation range of 140°. Diffraction data from a
C222 crystal were collected at beamline 4.2.2 of the Advanced Photon Source (900 frames, 180°
rotation). The data from both beamlines were integrated and scaled with XDS [16], and
intensities were merged and converted to amplitudes with AIMLESS [17]. The unit cell
dimensions of both crystal forms are listed in Table 1. The method of Matthews [18] predicts
eight chains in the P1 asymmetric unit with 49% solvent, and 2 chains in the C222 asymmetric
unit with 49% solvent. Data processing statistics are listed in Table 1.
Initial phases for the P1 structure were calculated using molecular replacement as
10

implemented MOLREP [19]. The search model was derived from a protomer of cod liver betaine
ALDH ([8], PDB ID: 1A4S, 70% identity to ALDH9A1). The side chains of the search model
were trimmed to the C-γ atom with CHAINSAW [20]. MOLREP returned a solution consisting
of eight protein chains arranged as two tetramers. We note that the molecular replacement
solutions were not as clear when search models derived from the structure of apo ALDH9A1
(PDB ID: 6QAP) were used, indicating conformational differences between our structure and
apo ALDH9A1.
Initial phases for the C222 structure were calculated using molecular replacement with
Phaser [21]. The search model was derived from a partial structure obtained by refinement
against an earlier C222 data set, which exhibited pseudo-merohedral twinning. Initial phases for
the twinned data set were calculated using Phaser with a search model derived from cod liver
betaine ALDH (PDB ID: 1A4S).
PHENIX [22] was used for refinement, and Coot [23] was used for model building. The B-
factor model consisted of an isotropic B-factor for each non-hydrogen atoms and one TLS group
per chain. Because of the modest resolution, non-crystallographic symmetry restraints were
+
enforced during refinement of both structures. The occupancies of the NAD molecules in both
structures were set to 1.0. The structures were validated with MolProbity [24] and the wwPDB
validation service [25]. Refinement statistics are listed in Table 1.
2
.7. Analytical ultracentrifugation
Sedimentation velocity experiments were performed in a Beckman XL-I analytical
ultracentrifuge using an An50Ti rotor at 20°C. For this analysis, His-tagged ALDH9A1 at 3 mg
-1
mL protein sample was dialyzed overnight against a buffer containing 50 mM HEPES pH 7.8,
11

1
00 mM NaCl, 5% (v/v) glycerol, and 0.5 mM DTT. A two-sector charcoal-Epon sedimentation
velocity centerpiece was loaded with reference buffer and protein samples. After an equilibration
period of 2 h, the sample was centrifuged at 35,000 rpm for a total of 300 radial scans spaced at 2
min intervals. Data were acquired using Rayleigh interference optics, which allowed
-1
measurement of the actual experimental protein concentration, assuming 1 mg mL is equivalent
to 3.33 fringes [26]. Scans 10 – 300 were analyzed and the distribution of apparent sedimentation
coefficients, c(s), and distribution of apparent molecular masses, c(M), were determined using
SEDFIT [27].
2
.8. Small-angle X-ray scattering
Shutterless SAXS data collection was performed at beamline 12.3.1 of the Advanced Light
Source through the SIBYLS Mail-in High Throughput SAXS program [28]. Prior to SAXS
analysis, purified His-tag-free ALDH9A1 was passed over a Superdex 200 10-30 size-exclusion
chromatography column in the presence of a buffer containing 50 mM HEPES pH 8.0, 600 mM
+
NaCl, 2% (v/v) glycerol, 1 mM DTT, and 1 mM NAD . Samples were then supplemented with 1
+
mM NAD and dialyzed overnight at 4°C against a buffer containing 50 mM HEPES pH 8.0,
+
6
00 mM NaCl, 2% (v/v) glycerol, 1 mM DTT, and 1 mM NAD .
SAXS data were collected on a Pilatus detector operating in shutterless mode, writing frames
every 0.3 s. Buffer subtracted SAXS curves were averaged using SAXS FrameSlice. PRIMUS
29] was used to inspect the merged data and to derive SAXS parameters. The maximum particle
[
dimension was estimated from calculations of the pair distribution function using GNOM [30]
via PRIMUS. The molecular mass was estimated using the SAXSMoW [31]. Theoretical SAXS
curves were calculated using FoXS [32]. DENSSWeb [33] was used for ab initio electron
12

density determination from solution scattering data.
2
.9. Negative stain electron microscopy
Preparation of negative-stained grids was carried out adapted from the previously described
protocol [34]. Briefly, carbon-coated copper grids (Electron Microscopy Sciences, Hatfield, PA)
-1
were glow discharged, and a 5 µL drop of His-tagged ALDH9A1 (~0.08 mg mL ) was applied
to the grid, followed by a 2 min incubation at room temperature. Excess protein was removed by
blotting with Whatman P4 filter paper. Blotting was immediately followed by two quick washes
in water and a quick wash in 0.75% (w/v) uranyl formate (UF; Ted Pella, Reading, CA), blotting
with filter paper in between wash steps. A final incubation, by floating the grid for 2 min on a
0
.75% UF (w/v) drop, was performed before excess UF was removed from the grid with filter
paper. The grid was air-dried. Protein particles were observed using a JEOL JEM 1400
transmission electron microscope (Peabody, MA) at 80 kV.
The data were analyzed with RELION-3 [35]. Contrast transfer function correction was
performed using CTFFind 4.1 using a spherical aberration of 3.2 mm, amplitude contrast of 0.5,
and magnified pixel size of 3.51 Å. The resolution range used for fitting was 5–30 Å. The
defocus range was 5000–50000 Å with a defocus step of 500 Å and an astigmatism of 200 Å. An
initial particle set was obtained using template-free auto-picking (Laplacian-of-Gaussian-based)
with a minimum diameter of 50 Å, maximum diameter of 120 Å, and threshold of 3.5. This
calculation generated 10373 particles. Low quality particles (e.g., random noise, overlapped
protein molecules) were eliminated during several rounds of iterative 2D class averaging and
selection, resulting in a set of 7430 particles that was used for the final 2D class average
calculation presented here.
13

3
. Results
3
.1. Time-dependent inhibition of ALDH9A1 by DEAB
DEAB (Fig. 1A) is variously a substrate, competitive tight-binding inhibitor, or irreversible
covalent inactivator of ALDHs [10, 36]. We therefore performed activity assays to understand
how DEAB interacts with ALDH9A1.
We first tested whether DEAB is a substrate using the approach of Morgan et al. [10] by
+
incubating ALDH9A1 (0.5 µM) with DEAB (30 µM) and NAD (1.5 mM), while monitoring the
absorbance at 360 nm and 300 nm. In this assay, a decrease at 360 nm indicates consumption of
DEAB, while an increase at 300 nm indicates production of the carboxylic acid product,
diethylaminobenzoic acid. The spectrum of DEAB alone shows a broad peak centered at 360 nm,
+
as expected (Fig. 1A). Spectra acquired for the mixture of ALDH9A1, DEAB, and NAD show
little change over 4 hours (Fig. 1A). These results suggest that DEAB is not an efficient substrate
for ALDH9A1.
Because we observed no significant activity with DEAB as the aldehyde substrate, we next
performed time-dependent inhibition assays. A progress curve approach was used in which the
+
enzyme (0.5 µM) was mixed with the aldehyde substrate hexanal (2 mM), NAD (2.5 mM), and
DEAB (0 – 5.0 µM) (Fig. 1B). The initial increase in absorbance at 340 nm indicates the
production of NADH associated with the catalytic turnover of hexanal to hexanoic acid. The
flattening of the curves at longer time is characteristic of covalent inactivation, similar to what
has been observed with ALDH2 [10] and ALDH7A1 [36]. Alternatively, time-dependent loss of
activity is associated with slow, tight-binding inhibition [13, 37, 38]. A control experiment
confirmed that flattening of the curves was not due to depletion of substrate or cofactor (inset,
Fig. 1B). Global fitting of the data to combined Equations 1a and 1b was performed to estimate
14

kinetic parameters for enzyme inactivation, assuming k
6
5
<< k (see Material and Methods). These
-1
calculations yielded k of 0.184 ± 0.052 min , K of 0.054 ± 0.018 µM, and an apparent second-
5
I
-1 -1
order rate constant for enzyme inactivation by DEAB (k /K ) of 57000 ± 25000 M s . This
5
I
-1 -1
value is similar to that of ALDH2 (86000 M s ) [10].
The reversibility of inhibition was assessed by monitoring the return of enzyme activity upon
+
dilution. ALDH9A1 (5 µM) was incubated with 10 µM DEAB and 1 mM NAD at 4°C for 60
minutes, and then assayed for activity by diluting 10-fold into assay buffer containing 0.5 mM
+
hexanal and 1 mM NAD . The progress curve for the DEAB-treated sample shows a ~5-minute
lag in NADH production, followed by an increase in activity (Fig. 2A). The steady-state velocity
of the DEAB-treated sample was ~70% compared to a control that was treated identically but
lacked DEAB. These results show that inhibition of ALDH9A1 by DEAB is reversible.
The reversibility of inhibition was confirmed by measuring the return of enzyme activity
+
after removal of DEAB. ALDH9A1 (5 µM) was incubated with 10 µM DEAB and 1 mM NAD
for 60 minutes, and then the excess DEAB was removed by spin ultrafiltration. The activity was
+
then measured in an assay containing 0.5 mM hexanal and 1 mM NAD . The progress curves for
the DEAB-treated and control samples are very similar (Fig. 2B). Note the progress curve for the
DEAB-treated sample does not exhibit a lag. These results are consistent with a covalent
reversible mechanism of inhibition.
+
3
.2. The fold of ALDH9A1 complexed with NAD
+
Two structures of ALDH9A1 complexed with NAD were determined. A 2.5 Å resolution
structure with space group P1 was obtained using enzyme that had been incubated with both
+
NAD and DEAB (Table 1). Also, a 2.64 Å structure in space group C222 was determined using
15

+
enzyme incubated with NAD , without DEAB. We note these crystal forms are different from
those used for structure determination of apo ALDH9A1 [7].
+
As expected, ALDH9A1 complexed with NAD exhibits the 3-domain architecture of apo
+
ALDH9A1, as well as other ALDHs (Fig. 3A). The NAD -binding domain features a 5-stranded
Rossmann-dinucleotide binding fold at its core. The catalytic domain has an α/β fold and
contains the catalytic cysteine residue (Cys288). The oligomerization domain adopts the
+
expected β- substructure protruding from the NAD -binding domain.
+
The structures of ALDH9A1-NAD were compared to each other and with the contents of
the PDB using PDBeFold [39] to identify the closest structural neighbors. Table 2 lists the
pairwise RMSD values between the chains of the various structures used in the analysis.
+
Comparison of our P1 and C222 NAD structures yields RMSD values of 1.0 – 1.3 Å. This
range is 3-5 times larger than the chain-chain RMSDs within either structure, suggesting that the
two structures represent distinct conformations. This idea was supported by comparing our
+
structures to the PDB. We found that the P1 ALDH9A1-NAD structure is most similar to
+
betaine ALDH from cod liver (RMSD = 0.6 – 0.7), whereas the C222 ALDH9A1-NAD
structure most resembles apo ALDH9A1 (RMSD = 0.4 – 0.6 Å).
The RMSD data suggest that the current ALDH9A1 structures can be classified into two
+
groups, one corresponding to the P1 NAD complex structure, and the other consisting of the
+
C222 NAD complex structure and the apo structures. Additional analysis shows that the two
+
groups differ by a domain rotation. For example, when the NAD -binding domains of the P1
+
NAD complex and apo ALDH9A1 are superimposed, the catalytic domains differ by an
apparent rigid body rotation of ~5° (Fig. 3B). Apparently, interactions of the enzyme with DEAB
+
+
and NAD caused the NAD -binding and catalytic domains to rotate closer together.
16

+
3
.3. Conformation and interactions of NAD bound to ALDH9A1
+
Electron density for NAD was observed in all eight chains of the P1 asymmetric unit. In
chain A, electron density was present for the entire cofactor (Fig. 4A), whereas in the other
chains only the ADP portion could be modeled with confidence (Fig. S3 of Supplementary data).
Similarly, only the AMP fragment could be modeled in the C222 structure (Fig. S4 of
+
Supplementary data). We note that weak density for the nicotinamide riboside of NAD is
common in ALDHs [40-44]. Electron density extending from catalytic Cys288 was present in all
eight chains of the P1 structure, possibly indicating covalent modification by DEAB; however,
we were not able to model DEAB covalently bound to Cys288 satisfactorily into these features.
The lack of convincing density for the covalent adduct is perhaps consistent with the apparent
reversibility of inactivation (Fig. 2).
+
NAD binds in the expected site at the C-termini of the β-strands of the Rossmann fold.
+
NAD forms several electrostatic interactions with the protein (Fig. 4A). The adenine ribose
hydrogen bonds with Lys180. The pyrophosphate interacts with Trp156, Ser233, and Thr236.
+
The nicotinamide ribose of the one complete NAD forms a hydrogen bond with Glu391, a
residue identically conserved in the ALDH superfamily [45].
+
The conformation of the complete NAD (chain A of P1 structure) is unusual in that the
nicotinamide riboside group is shifted by 3 – 4 Å out of the active site compared to the normal
(a.k.a. hydride transfer) conformation (Fig. 4B). For example, a structure of betaine ALDH
shows the hydride transfer conformation (Fig. 4B). The retracted pose in our structure is similar
to the hydrolysis conformation described by the Hurley group in their studies of ALDH2 [40]
(
Fig. 4B). A key difference between the hydride transfer and hydrolysis conformations is that the
17

former makes two hydrogen bonds to a conserved glutamate residue, whereas the latter makes
+
only one (Fig. 4B). Note the NAD in our structure exhibits this feature of the hydrolysis
conformation.
3
.4. Conformations of the
αE-βE region of the Rossmann fold and the inter-domain linker
The αE-βE region of the Rossmann fold (residues 232 – 258) exhibits a well-defined
+
+
conformation in the P1 NAD complex, in contrast to the C222 NAD -complex and apo enzyme
structures in which these residues are disordered. The electron density is strong and continuous
for the main chain, and most of the side chains also have clear electron density indicating their
conformations (Fig. 5A). Residues 234 – 247 form an α-helix, and residues 251 – 254 form a β-
strand (Fig. 5A); these are the final two secondary structural elements of the Rossmann
dinucleotide binding fold. Comparison to betaine ALDH shows that the αE-βE region of P1
+
ALDH9A1-NAD adopts the canonical secondary and tertiary structure of the ALDH
superfamily (Fig. 5B). Furthermore, the αE helices of the domain-swapped dimer interact across
a molecular two-fold axis, as in other ALDHs (Fig. 5C).
The electron density maps also clearly defined the conformation of the inter-domain linker in
+
the P1 NAD complex (residues 451 – 470) (Fig. 6A). This section of the polypeptide chain
connects the catalytic and oligomerization domains, and forms part of the active site. The linker
in our P1 structure adopts the β-hairpin structure typical of the ALDH superfamily, as shown by
the comparison with betaine ALDH (Fig. 6B). Several interactions within the β-hairpin stabilize
its conformation, including three main-chain hydrogen bonds and charged-hydrogen bonds
between Arg467 and the carbonyls of Gly466 and Gly459. These interactions are also present in
betaine ALDH. The canonical β-hairpin is very different from the S-shaped conformation in apo
18

ALDH9A1 (Fig. 6C). We note that the electron density for the inter-domain linker in the C222
+
NAD complex is weak and discontinuous, but nevertheless consistent with the S-conformation.
The ordering of αE-βE and the remodeling of the inter-domain linker result in the formation
of new tertiary and quaternary structural interactions (Fig. 7). For example, βE makes three main
chain hydrogen bonds with the inter-domain linker to form a two-stranded anti-parallel β-sheet
structure (Fig. 7A). And the tip of the inter-domain linker forms several hydrogen bonds with the
catalytic domain. The αE helices of adjacent protomers pack together to form a new dimer
interface (Fig. 7B). This interface involves the nonpolar face of αE, which includes Met238,
Met241, Ala245, and Ile248. Also, Phe465 of the linker intercalates between the βE strands of
two adjacent protomers, forming intermolecular nonpolar contacts with Ile248 and Pro250 (Fig.
7
B). In summary, the isomerization of the αE-βE region and the inter-domain linker into their
canonical conformations results in the formation of many noncovalent interactions that stabilize
the active form of ALDH9A1.
3
.5. Oligomeric structure of ALDH9A1
Sedimentation velocity analytical ultracentrifugation was used to determine the oligomeric
state of ALDH9A1 in solution. The distribution of apparent sedimentation coefficients, c(s), of
+
-1
apo (NAD -free) His-ALDH9A1 at 3 mg mL revealed a single major peak at 6.8 S (Fig. 8).
Likewise, the distribution of molecular masses, c(M), revealed a single major peak
corresponding to a molecular mass of 223 kDa, which is within 1% of the molecular mass of the
His-ALDH9A1 tetramer (theoretical molecular mass: 224.5 kDa). Thus, sedimentation velocity
-1
suggests apo His-ALDH9A1 is tetrameric in solution at 3 mg mL . This result is consistent with
the observation of the classic dimer-of-dimers ALDH tetramer in both of our structures, as well
19

as the crystal forms reported by Koncitikova et al. [7].
The in-solution quaternary structure of ALDH9A1 was determined using SAXS. Static
-1
SAXS data were collected at three nominal protein concentrations in the range of 1 – 5 mg mL ,
+
in the presence of 1 mM NAD (Fig. 9A) The average R from Guinier analysis of 37.9 ± 0.3 Å
g
(Table 3) is in agreement with the SAXS R
g
of 38 Å for the ALDH7A1 tetramer [43, 46] and the
R
g
of 35.5 Å calculated from the ALDH9A1 crystallographic tetramer. For each concentration,
the theoretical SAXS curve generated from the crystallographic tetramer provided a reasonably
good fit to the experimental data (Fig. 9A). Imposition of a multi-body fit to simulate a mixture
of dimer and tetramer did not statistically improve the fits to the experimental data. Finally, the
ab initio shape reconstruction shows good agreement with the tetramer (Fig. 9B). Overall, these
data suggest ALDH9A1 is primarily tetrameric under the conditions used, consistent with the
sedimentation velocity data.
The quaternary structure of apo ALDH9A1 in solution was confirmed with negative stain
electron microscopy (Fig. 10). The micrographs exhibited good contrast and particle separation
(
Fig. 10A). The 2D class averages calculated from 7430 particles are shown in Fig. 10B, and for
comparison, views of the ALDH9A1 tetramer and dimer are shown in Fig. 10C. Most of the 2D
classes appear to resemble the tetramer viewed down one of its three mutually orthogonal 2-fold
axes (classes 1, 4, 6-10). In these classes, which account for ~80% of the particles, the four
subunits of the tetramer and a two-fold symmetry axis perpendicular to the page are apparent.
Classes 3 and 5 (16% of the particles) appear to represent the tetramer rotated by 45° around a 2-
fold axis. Class 2 may represent a dimer viewed down its 2-fold axis. Note the dimer represents
only 4% of the particles. Thus, apo ALDH9A1 is predominantly tetrameric under the conditions
used for negative stain EM. In summary, the conclusion from multiple biophysical measurements
20

–
X-ray crystallography, sedimentation velocity, SAXS, and electron microscopy – is that
ALDH9A1 exists in solution primarily as the classic ALDH superfamily dimer-of-dimers
tetramer under the conditions used.
4
. Discussion
The new structures described here provide insight into the mechanism by which ALDH9A1
is activated for catalysis. The inactive conformation first observed in the apo enzyme is
characterized by disorder of αE-βE and a novel S-shaped inter-domain linker [7]. Because these
+
+
atypical features are also present in our C222 NAD -complex structure, it appears that NAD -
binding is not sufficient to activate the enzyme. Instead, we found that treatment of ALDH9A1
+
with DEAB and the cofactor NAD apparently induced αE-βE and the inter-domain linker to
assume their canonical conformations, as observed in the P1 structure. We conclude that the
+
presence of an aldehyde substrate and NAD promotes isomerization of the enzyme into the
active conformation.
The disorder-order transitions discussed here are not unique to ALDH9A1. Disorder in the
interdomain linker has also been observed in ALDH1A2 (PDB ID: 1BI9) [47], ALDH4A1
(PDB IDs: 4OE5/6) [42], and ALDH7A1 (PDB IDs: 4ZVX/Y) [43]. In ALDH1A2, ordering of
the linker is thought to be related to aldehyde binding, similar to ALDH9A1 [48]. In ALDH4A1
and ALDH7A1, no clear connection between ligand binding and disorder-order transitions has
been established. Disorder in αE-βE is rarer. The best example is a naturally-occurring disease
variant of ALDH2 (ALDH2*2) in which Glu487 is mutated to Lys (PDB ID: 1ZUM) [49].
Glu487 in wild-type ALDH2 forms intersubunit ion pairs with two arginine residues, one located
on βE (Arg264), and another in the interdomain linker (Arg475). The introduction of Lys487
21

obviously disrupts these stabilizing interactions, resulting in substantial disorder in both αE-βE
and the interdomain linker. ALDH9A1 does not have an analogous ionic interaction network,
due to the substitution of a glutamine for Glu487, and perhaps more importantly, a proline in
place of Arg264. The absence of these interactions may explain the propensity for disorder in
ALDH9A1. On the other hand, the close ALDH9A1 homolog, betaine ALDH, also lacks the
cross-subunit ion pairs, yet it exhibits canonical αE-βE and interdomain linker conformations,
even in the absence of bound ligands (PDB ID: 1A4S). Although the mechanisms of structural
ordering in ALDHs remain to be determined, the observation of large swaths of disorder in the
active sites of multiple ALDHs suggests disorder-order transitions may be an inherent feature of
the ALDH superfamily fold and important for catalytic function.
+
The NAD in our P1 structure unexpectedly has a retracted pose (Fig. 4). Two conformations
of the cofactor bound to ALDHs have been described: “hydride transfer” and “hydrolysis”.
+
These designations refer to steps in the catalytic mechanism. NAD tends to bind in the hydride
+
transfer conformation, which positions the hydride acceptor of NAD (C4 atom) ~3 Å from the
S-atom of the catalytic cysteine, ready to accept a hydride ion from the thiohemiacetal
intermediate. Following hydride transfer and formation of the acyl-enzyme intermediate, the
newly formed NADH retracts from the active site into the hydrolysis conformation. This
movement allows room for a water molecule to enter the active site for hydrolysis of the acyl-
+
enzyme intermediate. NAD in crystal structures typically adopts the hydride transfer pose;
+
however, NAD has also been observed in the hydrolysis pose in ALDH1 (PDB ID: 1BXS) [50]
and ALDH2 (PDB ID: 1O00) [40]. Also, mutation of the conserved glutamate that hydrogen
+
bonds to the nicotinamide ribose in ALDH7A1 causes NAD to switch to a hydrolysis-like pose
(PDB ID: 6O4L) [51]. NADH bound to ALDH2 adopts the mechanistically-expected hydrolysis
22

pose (PDB ID: 1O02) [40]. On the other hand, a structure of ALDH7A1 shows NADH in the
hydride transfer pose (PDB ID: 2J6L) [52]. Altogether, the crystal structures of ALDHs provide
+
evidence of the flexibility of the bound cofactor, which is essential for catalysis. Why NAD in
our structure is retracted into a hydrolysis-like conformation is unclear. It is possible that
covalent modification of the catalytic cysteine by DEAB, which is suggested by the electron
+
density but could not be modeled with certainty, prevents NAD from moving into the hydride
transfer conformation.
We showed that ALDH9A1 forms the classic ALDH superfamily dimer-of-dimers tetramer
in solution. This is not surprising considering that the closest sequence homologs of ALDH9A1
in the PDB also form the tetramer (at least in crystallo), including betaine ALDHs (40 – 70%
identical [8, 53]) and ALDH2 (43% identical).
ALDH9A1 may be added to the list of ALDHs that are inhibited by DEAB. This information
is pertinent to studies that use the ALDEFLUOR flow cytometry assay to detect cancer stem
cells based on ALDH activity (ALDH-bright cells) [54-58]. DEAB is used in the control arm of
the ALDEFLUOR assay, and any ALDHs that are reversibly inhibited or covalently inactivated
by DEAB will contribute to a cell being labeled as ALDH-bright and thus classified as a cancer
stem cell. DEAB was once thought to be specific for ALDH1A1, which led researchers to
assume that the ALDEFLUOR assay was specific for the detection of ALDH1A1. Recent work
has proven this assumption incorrect, as DEAB inhibits or inactivates several ALDHs including
ALDH1A1, ALDH1A3, ALDH1B1, ALDH5A1, ALDH1A2, ALDH2, and ALDH7A1 [10, 36,
5
9].
Hurley’s group proposed a general mechanism for the inhibition of ALDHs by DEAB. They
proposed that DEAB functions as a substrate through the hydride transfer step, but that the acyl-
23

enzyme intermediate is protected against hydrolysis by resonance stabilization [10]. The extent
of resonance stabilization, and therefore the potency of inhibition, depends on the noncovalent
interactions between the acyl-enzyme and the active site, which varies among ALDH isozymes.
For example, the active sites of ALDH1A2, ALDH2 and ALDH7A1 apparently provide
exceptional stabilization, since DEAB is a covalent irreversible inhibitor of these enzymes [10,
3
6]. For ALDH1A1, ALDH1A3, ALDH1B1, and ALDH5A1, DEAB inhibits by virtue of being
a slow substrate; the active sites of these enzymes apparently provide less efficient stabilization
of the acyl-enzyme than in ALDH1A2, ALDH2 and ALDH7A1.
We found that the inhibition of ALDH9A1 by DEAB is time-dependent, consistent with
covalent inactivation. In contrast to ALDH2 and ALDH7A1, the inhibition was diminished by
jump-dilution and abrogated by removal of excess DEAB, consistent with a covalent, reversible
mechanism. The irreversible inactivation of ALDH2 and ALDH7A1 by DEAB in the presence of
+
NAD is due to formation of the acyl-enzyme [10, 36]. In principle, covalent reversible
inhibition of an ALDH can arise by slow hydrolysis of the acyl-enzyme intermediate to generate
active enzyme and diethylaminobenzoic acid. This is not likely to be the case for ALDH9A1,
given the rapid return of enzyme activity upon removal of DEAB coupled with the fact that we
find no evidence that DEAB serves as a substrate for this enzyme (see Fig. 1A). A more
plausible mechanism is formation of a reversible thiohemiacetal covalent intermediate (Scheme
2
A) that fails to progress through the hydride transfer step of the normal catalytic mechanism
that would generate the acyl-enzyme intermediate. The covalent, reversible inhibition due to
thiohemiacetal formation with an active site cysteine residue proposed here finds a direct
mechanistic analogy in the inhibition cysteine proteases by aldehydes [60-62] and is
mechanistically related to the inhibition of viral proteases by ketoamide compounds that form a
24

reversible, covalent tetrahedral species with the catalytic serine [63-65]. This mechanism
(Scheme 2A) could also explain the covalent, reversible inhibition of ALDH1A2 by DEAB in
+
the absence of NAD [10].
The progress curves in Figure 1B and the reversibility of inhibition are also consistent with
slow, tight-binding inhibition (Scheme 2B) [13]. In this mechanism, inhibitor binding could
involve the rapid formation of an initial collision complex (E⋅I) that subsequently undergoes a
slow isomerization reaction to E⋅I* (with or without formation of the thiohemiacetal covalent
intermediate). Presumably the initial encounter would involve the apo, inactive enzyme,
followed by an isomerization step resulting in an enzyme conformation similar to the P1
structure described here. Isomerization between these two conformations likely would be quite
slow (seconds to minutes), as it involves very large structural changes, including folding of the
αE-βE region and reorganization of the interdomain linker.
Funding
Research reported in this publication was supported by the NIGMS of the National Institutes of
Health under award number R01GM093123 (to J.J.T.), and an administrative supplement to NIH
grant R01GM065546 (to J.J.T., Collaborative Supplements for Cryo-Electron Microscopy
Technology Transfer). I.A.Q. was supported by an ICMR International Fellowship from the
Indian Council of Medical Research, India.
Acknowledgements
We thank K. Burnett for collecting the SAXS data through the SIBYLS mail-in program, J.
Schuermann for help with data collection at beamline 24-ID-E, J. Nix for help with data
25

collection at beamline 4.2.2, and T. White for help with EM data collection. This work is based
upon research conducted at the Northeastern Collaborative Access Team beamlines, which are
funded by the National Institute of General Medical Sciences from the National Institutes of
Health (P30 GM124165). The Eiger 16M detector on 24-ID-E beam line is funded by a NIH-
ORIP HEI grant (S10OD021527). This research used resources of the Advanced Photon Source,
a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE
Office of Science by Argonne National Laboratory under Contract No. DE-AC02-06CH11357.
Part of this work was conducted at the Advanced Light Source (ALS), a national user facility
operated by Lawrence Berkeley National Laboratory on behalf of the Department of Energy,
Office of Basic Energy Sciences, through the Integrated Diffraction Analysis Technologies
(
IDAT) program, supported by DOE Office of Biological and Environmental Research.
Additional support comes from the National Institute of Health project ALS-ENABLE (P30
GM124169) and a High-End Instrumentation Grant S10OD018483.
References
[
1] N.A. Sophos, V. Vasiliou, Aldehyde dehydrogenase gene superfamily: the 2002 update,
Chem. Biol .Interact. 143-144 (2003) 5-22.
2] V. Koppaka, D.C. Thompson, Y. Chen, M. Ellermann, K.C. Nicolaou, R.O. Juvonen, D.
[
Petersen, R.A. Deitrich, T.D. Hurley, V. Vasiliou, Aldehyde dehydrogenase inhibitors: a
comprehensive review of the pharmacology, mechanism of action, substrate specificity, and
clinical application, Pharmacol. Rev. 64(3) (2012) 520-39.
[3] F.M. Vaz, S.W. Fouchier, R. Ofman, M. Sommer, R.J. Wanders, Molecular and biochemical
characterization of rat gamma-trimethylaminobutyraldehyde dehydrogenase and evidence for the
involvement of human aldehyde dehydrogenase 9 in carnitine biosynthesis, J Biol Chem 275(10)
(2000) 7390-4.
[4] G. Kurys, W. Ambroziak, R. Pietruszko, Human aldehyde dehydrogenase. Purification and
characterization of a third isozyme with low Km for gamma-aminobutyraldehyde, J Biol Chem
64(8) (1989) 4715-21.
5] W. Ambroziak, R. Pietruszko, Human aldehyde dehydrogenase. Activity with aldehyde
metabolites of monoamines, diamines, and polyamines, J Biol Chem 266(20) (1991) 13011-8.
2
[
26

[
6] M.K. Chern, R. Pietruszko, Human aldehyde dehydrogenase E3 isozyme is a betaine
aldehyde dehydrogenase, Biochemical and biophysical research communications 213(2) (1995)
61-8.
7] R. Koncitikova, A. Vigouroux, M. Kopecna, M. Sebela, S. Morera, D. Kopecny, Kinetic and
structural analysis of human ALDH9A1, Bioscience reports 39(4) (2019).
8] K. Johansson, M. El-Ahmad, S. Ramaswamy, L. Hjelmqvist, H. Jornvall, H. Eklund,
Structure of betaine aldehyde dehydrogenase at 2.1 A resolution, Protein Sci 7(10) (1998) 2106-
7.
9] E. Gasteiger, C. Hoogland, A. Gattiker, S. Duvaud, M.R. Wilkins, R.D. Appel, A. Bairoch,
Protein identification and analysis tools on ExPASy server, Humana Press, Totowa, NJ, 2005.
10] C.A. Morgan, B. Parajuli, C.D. Buchman, K. Dria, T.D. Hurley, N,N-
5
[
[
1
[
[
diethylaminobenzaldehyde (DEAB) as a substrate and mechanism-based inhibitor for human
ALDH isoenzymes, Chem Biol Interact 234 (2015) 18-28.
[
11] J.I. Juncosa, K. Takaya, H.V. Le, M.J. Moschitto, P.M. Weerawarna, R. Mascarenhas, D.
Liu, S.L. Dewey, R.B. Silverman, Design and Mechanism of (S)-3-Amino-4-
difluoromethylenyl)cyclopent-1-ene-1-carboxylic Acid, a Highly Potent gamma-Aminobutyric
Acid Aminotransferase Inactivator for the Treatment of Addiction, J Am Chem Soc 140(6)
(
(2018) 2151-2164.
[12] K.A. Salminen, J. Leppanen, J.I. Venalainen, M. Pasanen, S. Auriola, R.O. Juvonen, H.
Raunio, Simple, direct, and informative method for the assessment of CYP2C19 enzyme
inactivation kinetics, Drug Metab Dispos 39(3) (2011) 412-8.
[
13] J.F. Morrison, C.T. Walsh, The behavior and significance of slow-binding enzyme
inhibitors, Adv Enzymol Relat Areas Mol Biol 61 (1988) 201-301.
14] A. D'Arcy, T. Bergfors, S.W. Cowan-Jacob, M. Marsh, Microseed matrix screening for
[
optimization in protein crystallization: what have we learned?, Acta Crystallogr F Struct Biol
Commun 70(Pt 9) (2014) 1117-26.
[15] M. Till, A. Robson, M.J. Byrne, A.V. Nair, S.A. Kolek, P.D. Shaw Stewart, P.R. Race,
Improving the success rate of protein crystallization by random microseed matrix screening, J
Vis Exp (78) (2013).
[
[
16] W. Kabsch, XDS, Acta Crystallogr D Biol Crystallogr 66(Pt 2) (2010) 125-32.
17] P.R. Evans, G.N. Murshudov, How good are my data and what is the resolution?, Acta
Crystallogr. D Biol. Crystallogr. 69(Pt 7) (2013) 1204-14.
[
[
18] B.W. Matthews, Solvent content of protein crystals, J. Mol. Biol. 33 (1968) 491-497.
19] A. Vagin, A. Teplyakov, MOLREP: an automated program for molecular replacement, J.
Appl. Crystallogr. 30 (1997) 1022-1025.
20] N. Stein, CHAINSAW: a program for mutating pdb files used as templates in molecular
replacement, J. Appl. Crystallogr. 41(3) (2008) 641-643.
21] A.J. McCoy, R.W. Grosse-Kunstleve, P.D. Adams, M.D. Winn, L.C. Storoni, R.J. Read,
Phaser crystallographic software, J. Appl. Crystallogr. 40(Pt 4) (2007) 658-674.
22] P.V. Afonine, R.W. Grosse-Kunstleve, N. Echols, J.J. Headd, N.W. Moriarty, M.
[
[
[
Mustyakimov, T.C. Terwilliger, A. Urzhumtsev, P.H. Zwart, P.D. Adams, Towards automated
crystallographic structure refinement with phenix.refine, Acta Crystallogr. D Biol. Crystallogr.
6
[
8(Pt 4) (2012) 352-67.
23] P. Emsley, B. Lohkamp, W.G. Scott, K. Cowtan, Features and development of Coot, Acta
Crystallogr D Biol Crystallogr 66(Pt 4) (2010) 486-501.
27

[24] V.B. Chen, W.B. Arendall, 3rd, J.J. Headd, D.A. Keedy, R.M. Immormino, G.J. Kapral,
L.W. Murray, J.S. Richardson, D.C. Richardson, MolProbity: all-atom structure validation for
macromolecular crystallography, Acta Crystallogr. D Biol. Crystallogr. D66(Pt 1) (2010) 12-21.
[25] S. Gore, E. Sanz Garcia, P.M.S. Hendrickx, A. Gutmanas, J.D. Westbrook, H. Yang, Z.
Feng, K. Baskaran, J.M. Berrisford, B.P. Hudson, Y. Ikegawa, N. Kobayashi, C.L. Lawson, S.
Mading, L. Mak, A. Mukhopadhyay, T.J. Oldfield, A. Patwardhan, E. Peisach, G. Sahni, M.R.
Sekharan, S. Sen, C. Shao, O.S. Smart, E.L. Ulrich, R. Yamashita, M. Quesada, J.Y. Young, H.
Nakamura, J.L. Markley, H.M. Berman, S.K. Burley, S. Velankar, G.J. Kleywegt, Validation of
Structures in the Protein Data Bank, Structure 25(12) (2017) 1916-1927.
[
26] J.L. Cole, J.W. Lary, P.M. T, T.M. Laue, Analytical ultracentrifugation: sedimentation
velocity and sedimentation equilibrium, Methods Cell Biol 84 (2008) 143-79.
27] P. Schuck, Size-distribution analysis of macromolecules by sedimentation velocity
ultracentrifugation and lamm equation modeling, Biophys. J. 78(3) (2000) 1606-19.
28] K.N. Dyer, M. Hammel, R.P. Rambo, S.E. Tsutakawa, I. Rodic, S. Classen, J.A. Tainer,
[
[
G.L. Hura, High-throughput SAXS for the characterization of biomolecules in solution: a
practical approach, Methods Mol Biol 1091 (2014) 245-58.
[29] P.V. Konarev, V.V. Volkov, A.V. Sokolova, M.H.J. Koch, D.I. Svergun, IUCr, PRIMUS: a
Windows PC-based system for small-angle scattering data analysis, Journal of applied
crystallography 36(5) (2003) 1277-1282.
[
30] D.I. Svergun, IUCr, Determination of the regularization parameter in indirect-transform
methods using perceptual criteria, Journal of applied crystallography 25(4) (1992) 495-503.
31] V. Piiadov, E. Ares de Araujo, M. Oliveira Neto, A.F. Craievich, I. Polikarpov, SAXSMoW
.0: Online calculator of the molecular weight of proteins in dilute solution from experimental
SAXS data measured on a relative scale, Protein Sci 28(2) (2019) 454-463.
32] D. Schneidman-Duhovny, M. Hammel, J.A. Tainer, A. Sali, FoXS, FoXSDock and
[
2
[
MultiFoXS: Single-state and multi-state structural modeling of proteins and their complexes
based on SAXS profiles., Nucleic Acids Res. 44(W1) (2016) W424-9.
[
33] T.D. Grant, Ab initio electron density determination directly from solution scattering data,
Nat Methods 15(3) (2018) 191-193.
34] M. Rames, Y. Yu, G. Ren, Optimized negative staining: a high-throughput protocol for
[
examining small and asymmetric protein structure by electron microscopy, J Vis Exp (90) (2014)
e51087.
[35] J. Zivanov, T. Nakane, B.O. Forsberg, D. Kimanius, W.J. Hagen, E. Lindahl, S.H. Scheres,
New tools for automated high-resolution cryo-EM structure determination in RELION-3, Elife 7
(2018).
[36] M. Luo, K.S. Gates, M.T. Henzl, J.J. Tanner, Diethylaminobenzaldehyde is a covalent,
irreversible inactivator of ALDH7A1, ACS Chem. Biol. 10(3) (2015) 693-7.
37] J. Pandhare, C. Dash, M. Rao, V. Deshpande, Slow tight binding inhibition of proteinase K
[
by a proteinaceous inhibitor: conformational alterations responsible for conferring irreversibility
to the enzyme-inhibitor complex, J Biol Chem 278(49) (2003) 48735-44.
[38] C. Dash, V. Vathipadiekal, S.P. George, M. Rao, Slow-tight binding inhibition of xylanase
by an aspartic protease inhibitor: kinetic parameters and conformational changes that determine
the affinity and selectivity of the bifunctional nature of the inhibitor, J Biol Chem 277(20) (2002)
1
7978-86.
28