Design, synthesis, and ex vivo evaluation of a selective inhibitor for retinaldehyde dehydrogenase enzymes

Article abstract of DOI:10.1016/j.bmc.2018.10.009

The retinaldehyde dehydrogenase (RALDH) enzymes, RALDH1, RALDH2, and RALDH3, catalyze the irreversible oxidation of retinaldehyde to all-trans-retinoic acid (ATRA). Despite the importance of the RALDH enzymes in embryonic development, postnatal growth and differentiation, and in several disease states, there are no commercially available inhibitors that specifically target these isozymes. We report here the development and characterization of a small molecule inhibitor dichloro-all-trans-retinone (DAR) (Summers et al., 2017) that is an irreversible inhibitor of RALDH1, 2, and 3 that effectively inhibits RALDH1, 2, and 3 in the nanomolar range but has no inhibitory activity against mitochondrial ALDH2. These results provide support for the development of DAR as a specific ATRA synthesis inhibitor for a variety of experimental and clinical applications.

Full text of DOI:10.1016/j.bmc.2018.10.009

Accepted Manuscript
Design, Synthesis, and Ex Vivo Evaluation of a Selective Inhibitor for Retinal-
dehyde Dehydrogenase Enzymes
Angelica R. Harper, Anh T. Le, Timothy Mather, Anthony Burgett, William
Berry, Jody A. Summers
PII:
S0968-0896(18)31382-8
https://doi.org/10.1016/j.bmc.2018.10.009
BMC 14568
DOI:
Reference:
To appear in:
Bioorganic & Medicinal Chemistry
Received Date:
Revised Date:
Accepted Date:
2 August 2018
3 October 2018
12 October 2018
Please cite this article as: Harper, A.R., Le, A.T., Mather, T., Burgett, A., Berry, W., Summers, J.A., Design,
Synthesis, and Ex Vivo Evaluation of a Selective Inhibitor for Retinaldehyde Dehydrogenase Enzymes, Bioorganic
&
Medicinal Chemistry (2018), doi: https://doi.org/10.1016/j.bmc.2018.10.009
This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers
we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and
review of the resulting proof before it is published in its final form. Please note that during the production process
errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

Design, Synthesis, and Ex Vivo Evaluation of a Selective
Inhibitor for Retinaldehyde Dehydrogenase Enzymes.
a,1,2
b,2
c
b
a
Angelica R. Harper , Anh T. Le , Timothy Mather , Anthony Burgett , William Berry , Jody A.
a
Summers
a
Department of Cell Biology, University of Oklahoma Health Sciences Center, Oklahoma City,
Oklahoma, 73104, United States
b
Department of Chemistry and Biochemistry, University of Oklahoma, Norman, Oklahoma
7
3019, United States
c
Oklahoma Medical Research Foundation, Oklahoma City, Oklahoma 73104, United States
Corresponding Author(s)
Jody A. Summers; jody-summers@ouhsc.edu; 1-405-271-8001, ext. 46478
1

ABSTRACT
The retinaldehyde dehydrogenase (RALDH) enzymes, RALDH1, RALDH2, and
RALDH3, catalyze the irreversible oxidation of retinaldehyde to all-trans-retinoic acid (ATRA).
Despite the importance of the RALDH enzymes in embryonic development, postnatal growth
and differentiation, and in several disease states, there are no commercially available inhibitors
that specifically target these isozymes. We report here the development and characterization of a
1
small molecule inhibitor dichloro-all-trans-retinone (DAR) that is an irreversible inhibitor of
RALDH1, 2, and 3 that effectively inhibits RALDH1, 2, and 3 in the nanomolar range but has no
inhibitory activity against mitochondrial ALDH2. These results provide support for the
development of DAR as a specific ATRA synthesis inhibitor for a variety of experimental and
clinical applications.
2

1
. INTRODUCTION
+
The aldehyde dehydrogenase (ALDHs) superfamily catalyzes the NAD(P) dependent,
irreversible oxidation of aldehydes into their respective carboxylic acids. Currently, there are 19
2
-4
known ALDH protein coding genes and 3 pseudogenes present in the human genome. ALDH
proteins are involved in a wide variety of physiological processes including aldehyde
5
-8
detoxification, alcohol metabolism, development, vision, and neurotransmission.
Mutations
and polymorphisms in ALDH genes have also been associated with a number diseases such as
brain, breast, and lung cancer, late-onset Alzheimer’s disease, alcohol flushing syndrome,
4, 9-19
Sjögren-Larsson syndrome (SLS), and type II hyperprolinemia.
The retinaldehyde dehydrogenase (RALDH, also ALDH1A) subfamily consists of three
members, RALDH1 (ALDH1A1), RALDH2 (ALDH1A2), and RALDH3 (ALDH1A3), which
synthesize all-trans-retinoic acid (ATRA) from retinaldehyde. All RALDH isozymes can utilize
both all-trans-retinaldehyde and 9-cis-retinaldehyde as substrates; however, RALDH2 exhibits
2
0, 21
the highest specificity for all-trans-retinaldehyde.
RALDH proteins are essential for
regulating ATRA signaling pre- and postnatally. In embryonic development, RALDH1, 2, and 3
are differentially expressed in a spatial-temporal manner to mediate organogenesis of the eye,
2
, 22-24
brain, heart, kidney, lung, and reproductive organs.
Postnatally, RALDH isozymes have
2
5-27
much more limited roles, but are involved in spermatogenesis,
maintenance of lens
The RALDH subfamily has also been
implicated in a variety of diseases through the modulation of all-trans-retinaldehyde and ATRA
28, 29
30-31
transparency,
and stem cell differentiation.
32-36
levels. RALDH1 has been linked to obesity, inflammation, and Crohn’s disease,
while
37, 38
RALDH2 has been associated with abnormal ocular growth (myopia).
Changes in the
3

expression of all three RALDH proteins in tumors have been associated with poor cancer
39-44
prognoses.
Despite the involvement of various ALDH family members in numerous disease states,
pharmacological inhibitors have been developed for only 3 of the ALDH isozymes: ALDH1A1,
3
ALDH2, and ALDH3A1. Of these three, selective inhibitors have been developed for
ALDH1A1 (CM026, CM037) and ALDH3A1 (CB29) for the treatment of oxazaphosphorine
45-47
resistant cancers.
Other inhibitors have been found to be effective against the above ALDHs;
3
however, these compounds are non-specific and inhibit a range of ALDH family members.
Non-specific ALDH inhibitors that have been effective at inhibiting the RALDH subfamily
include 4-amino-4methyl-2pentyne-1-al (AMPAL), citral, 4-(diethylamino)benzaldehyde
3
(
DEAB), disulfiram (commercially known as Antabuse), and gossypol. DEAB is an irreversible
inhibitor of ALDH1A2, ALDH2 and ALDH7A1 and has been utilized in studies of ALDH-
48-51
mediated drug metabolism and in the treatment of cancer.
Citral and gossypol are reversible
noncompetitive inhibitors of ALDH1, ALDH2, and ALDH3 isozymes and have been used in
5
2-56
experimental studies on embryonic development and cancer, respectively.
AMPAL and
disulfiram are irreversible inhibitors of ALDH1, ALDH2, and ALDH3 isoenzymes. AMPAL
5
7, 58
has been used to inhibit cell growth in tumors,
while disulfiram has been used commercially
59, 60
for the treatment of alcoholism, as an alcohol aversive agent.
Through inhibition of ALDH2,
disulfiram administration before drinking alcoholic beverages results in unpleasant symptoms
such as blurred vision, nausea, and flushing of the face and neck that are collectively known as
61-63
the disulfiram ethanol reaction.
Due to the lack of specificity of the aforementioned
inhibitors, studies utilizing these non-specific inhibitors to gain insights into the physiologic
roles of ALDH isoenzyme activity may be complicated by inhibition of multiple isoenzymes and
6
4-66
lead to off-target effects.
4

Recently, the bisdiamine, WIN 18446 (commercially known as Fertilysin), has been
6
7, 68
identified as a potent, irreversible RALDH2 inhibitor.
WIN 18446 has been used as a male
6
7, 69-71
contraceptive in a number of mammals
and was tested for its potential as a male
63
contraceptive in human volunteers. In all of these studies, WIN 18446 was found to effectively
and reversibly inhibit spermatogenesis, presumably through inhibition of RALDH2– mediated
2
4, 63, 72
testicular ATRA synthesis, recently shown to be required for normal spermatogenesis.
Although WIN 18446 was a promising male contraceptive, it was not marketed to the general
public, as the disulfiram ethanol reaction was observed upon patient consumption of alcohol,
indicating that WIN 18446 also inhibited ALDH2 and possibly other ALDH enzymes. Recently,
6
8
the non-specificity of WIN 18446 has been confirmed in vitro. These results demonstrate the
importance of developing small molecule inhibitors selective for the RALDH isozymes as such
inhibitors would have application for the study and treatment of a variety of diseases and
conditions. Using a structurally based drug design approach, we report here the development of
a novel RALDH (ALDH1A) selective inhibitor, (3E,5E,7E,9E)-1,1-dichloro-4,8-dimethyl-10-
(2,6,6-trimethylcyclohex-1-en-1-yl)deca-3,5,7,9-tetraen-2-one (“dichloro-all-trans-retinone” or
1
“
DAR”), that is an effective inhibitor of RALDH1 (IC = 434.70 ± 99.70 nM), RALDH2 (IC
5
0
50
=
55.00 ± 10.71 nM), and RALDH3 (IC = 161.27 ± 16.57 nM), but has no activity on human
5
0
mitochondrial ALDH2.
2
. RESULTS AND DISCUSSION
2
.1 STRUCTURE-BASED DESIGN OF DAR
WIN 18446 (Figure 1A) has been shown to inhibit RALDH1, 2, and 3, as well as
6
7, 68, 74
ALDH2 and other members of the ALDH superfamily.
Based on an x-ray crystal structure
of WIN 18446 bound to human RALDH2, the ALDH active cysteine sulfhydryl group forms a
5

covalent bond with the terminal carbon of WIN 18446, causing displacement of a chloride atom
7
5
from WIN 18446.
We postulated that the relatively large size of the substrate entrance tunnel
for the RALDH enzymes, as compared with the other ALDH enzymes, would allow for the
2
0, 76
specificity of retinaldehyde (RAL) (Figure 1B) to the RALDH enzymes.
Using this
knowledge, we suspected that a RALDH selective inhibitor could be generated by attaching a
dichloro-methane moiety to the retinyl group (beta-ionone ring with an isoprenoid chain) of
RAL, thereby forming dichloro-all-trans-retinone (DAR) (Figure 1C). Therefore, DAR (367.34
Da) was synthesized by reacting dichloromethyllithium with retinoic acid ethyl ester under
strong basic conditions. DAR has no structural similarity to commercially available aldehyde
dehydrogenase modulators or any small molecule in the CHemDiv, ZINC, or ChemSpider
databases.
[insert figure 1 here]
6

Scheme 1. One-Step Synthesis of α, α-Dichloro-Retinal Analog. Retinoic acid ethyl ester
was reacted with dichloromethyllithium generated in situ from dichloromethane (DCM) and
lithium diisopropylamide (LDA) in ethyl ether (Et O) to generate α,α-dichloro-all-trans-
2
retinone (DAR). 4 – 10% of DAR was produced in each reaction.
α,α-Dichloro-all-trans-retinone (“DAR”) was produced in a one-step transformation of
retinoic acid ethyl ester to DAR (Scheme 1). Dichloromethyllithium, generated in situ at low
7
3
temperature, reacted with commercially available retinoic acid ethyl ester to produce the α,α-
dichloro-retinoic acid in low yield (4 – 10%). A small amount of the starting material retinoic
acid ethyl ester was recovered, with the remaining mass balance of this reaction being comprised
of a complex mixture of unidentified isomeric byproducts. Extensive purification, including
semi-preparative HPLC, produced analytically pure DAR, free from any contaminating
byproducts. The low yield of this reaction is not surprising given the combination of the inherent
reactivity of the extended conjugated system in DAR and the highly reactive
dichloromethyllithium species.
2
.2. KINETIC CHARACTERIZATION OF DAR
RALDH activity in the presence and absence of DAR was measured by quantifying the
synthesis of NADH, as both NADH and ATRA are produced in equimolar amounts during the
3
oxidation of all-trans-retinaldehyde. Due to overlap in the absorption spectrum of RAL and
NADH (300 – 400 nm), a fluorescence-based NADH assay (AAT Bioquest, Inc.) was employed
to measure RALDH enzyme activity (ATRA synthesis) utilizing RAL as the substrate (Figure
2
). Additionally, ATRA was quantified by HPLC/spectrophotometric assay, and measurements
of ATRA synthesis were compared between the fluorescence based and
7

HPLC/spectrophotometric assays (Figure 2A). The fluorescence based assay exhibited high
correlation with HPLC measurements (p < 0.001, Pearson’s correlation analysis) when
recombinant chicken RALDH2 (2 μg) was incubated with RAL (0 – 100 μM) for 30 min.
Therefore the effects of protein concentration, time, and substrate concentration on NADH
synthesis, by chicken RALDH2 were examined (Figure 2B – D) as a proxy for ATRA synthesis.
NADH synthesis was linear between 0 – 2 μg RALDH2 (Figure 2B) and 0 – 50 min of
incubation time (Figure 2C); thus, all subsequent assays were conducted with 0.5 μg RALDH2
for 30 min, unless otherwise stated. The rate of NADH synthesis was saturated with increasing
concentrations of RAL (0 – 250 μM) (Figure 2D). Under these experimental conditions,
RALDH2 exhibited a K of 80.70 ± 20.38 μM for retinaldehyde (determined from four
m
independent experiments, each performed in triplicate). Previously reported Michaelis constants
for RALDH2 (K or K using retinaldehyde as the substrate) have ranged from 0.3 µM – 16
m
0.5
2
1, 67, 77, 78
µM.
method of assessing RALDH2 activity (HPLC quantification of ATRA, NADH absorbance at
40 nM, and NADH quantification using the fluorescence-based recycling system in the present
study).
Differences between these and our studies are most likely due to differences in the
3
[
insert figure 2 here]
To examine the effects of DAR on aldehyde dehydrogenase activity, DAR was tested on
recombinant chicken RALDH1, RALDH2, RALDH3, human RALDH2 (hRALDH2) and
recombinant human mitochondrial ALDH2 (hALDH2) (Figure 3, Table 1). RALDH1, 2, 3, and
hRALDH2 were pre-incubated with DAR (0 – 10 μM) for 20 min at 37°C, after which RAL (250
μM) was added to the reaction mixture to initiate ATRA and NADH production. DAR was
found to inhibit chicken RALDH1, 2, and 3 with IC ’s of 434.70 ± 99.70 nM, 55.00 ± 10.71
5
0
8

nM and 161.27 ± 16.57,nM, respectively, as well as human RALDH2 (hRALDH2; IC = 191.32
50
±
36.77) (Figure 3A – D, Table 1). Comparison of the IC values for DAR on RALDH1,
5
0
RALDH2, RALDH3 and hRALDH2 indicated significantly different inhibition for every pair of
enzymes (p < 0.05) with the exception of RALDH3 as compared with hRALDH2 (one way
ANOVA with Bonferroni correction for multiple comparisons).
The effect of DAR on
RALDH1 and RALDH3 activity demonstrated dose responses that were essentially step
functions, and thus data were fit using the 4-parameter logistics equation and the Hill-slope.
Using a spectrophotometric, absorbance assay measuring NADH production at 340 nm and
propionaldehyde as the substrate (5 mM), DAR(0 – 100 μM, 1 hr pre-incubation) did not have an
inhibitory effect on mitochondrial human aldehyde dehydrogenase 2 (hALDH2) (Figure 3E,
Table 1). As expected, hALDH2 was effectively inhibited by WIN 18446 (0 – 10 μM) with an
67
observed IC of 229.33 ± 40.95 nM, similar to that observed in other studies. These results
50
indicate that DAR has selectivity for the RALDH enzymes with the most robust inhibition
(
lowest IC ) observed for RALDH2 in vitro in the presence of the natural substrate, RAL
50
(inhibition of RALDH2 > RALDH3 > RALDH1).
Table 1. IC and K Values Determined for DAR
5
0
I
*
[
insert figure 3
here]
ALDH Isoenzyme
IC50
nM
K
I
(Morrison)
nM
RALDH1 (chick)
RALDH2 (chick)
RALDH3 (chick)
RALDH2 (human)
ALDH2 (human)
434.70 ± 99.70
55.00 ± 10.71
161.27 ± 16.57
191.32 ± 36.77
NI
194.40 ± 99.32
5.10 ± 2.50
6.74 ± 1.73
64.35 ± 10.76
NI
K ’s
were
Q=(Ki*(1+(S/Km))) and Y=Vo*(1-((((Et+X+Q)-(((Et+X+Q)^2)-
*Et*X)^0.5))/(2*Et))) (GraphPad Prism v5), where Et is the
determined
using
the
Morrison
equation:
I
4
concentration of enzyme (RALDH1 = 0.358 µM; RALDH2 = 0.091
µM; RALDH3 = 0.355 µM; hRALDH2 = 0.176 µM; ALDH2 = 4.43
µM), S is the concentration of substrate (described in Experimental
9
Section), and
Km is the Michaelis-Menten constant (see
supplemental Table S1). *Values are the mean ± SEM of triplicate
determinations from 3 ─ 5 independent experiments. NI = no
inhibition.

It is known that WIN 18446 inhibits human RALDH2 in a time-dependent and
6
8, 74, 79
irreversible manner.
To determine if DAR also acts as an irreversible inhibitor, the
mechanism of inhibition for recombinant chicken RALDH2 was investigated using various
methods (Figure 4). Increasing concentrations of DAR in the enzyme reaction resulted in a dose
dependent decrease of Vmax (1665, 1563, 1148, 866.9, 666.5, 503.0 pmol/μM RALDH2/min)
(Figure 4A, Table S1). A hallmark of irreversible inhibition is the persistence of inhibition
80, 81
following removal of unbound inhibitor.
When unbound inhibitor was removed by
ultrafiltration (Figure 4B; “UF”) prior to addition of cofactor and substrate, RALDH2 activity
was not restored. Significant inhibition was observed at 25 nM (33.02 ± 13.89% activity
remaining) and 50 nM (22.97 ± 4.55% activity remaining) of DAR compared to vehicle control
(100.00 ± 11.57% activity remaining) (p < 0.001, one-way ANOVA with Bonferroni’s test for
multiple comparisons), similar to the results seen when unbound DAR remained present in the
enzyme reaction (Figure 4B; “No UF”). In samples subjected to ultrafiltration, the average
inhibition of RALDH2 by DAR was greater than predicted by stoichiometric irreversible
1
0

inhibition. As can be observed in Figure 4B, the variability of ultracentrifuged (UF) samples was
- 4x higher than that of samples not ultracentrifuged (No UF) prior to enzyme assays. This
2
increased inhibition and variability is most likely due to variable loss of enzyme during recovery
from the centrifugal filter and/or variable loss of activity as a consequence of ultracentrifugation.
In addition, the Vmax for NADH synthesis was determined using increasing concentrations of
RALDH2 (46 – 365 nM) in the presence of DAR(150 nM) (Figure 4C). A plot of Vmax vs.
RALDH2 concentration indicated that RALDH2 activity was completely inhibited when enzyme
concentrations (46 – 91 nM) were less than the concentration of DAR ([RALDH2] ≤ 150 nM).
In the presence of 150 nM DAR, RALDH2 concentrations (183 – 365 nM) resulted in linear
increases in Vmax for NADH synthesis (0.50 – 3.01 μM NADH/min). To further characterize the
inhibitory effects of DAR, RALDH2 (0.5 µg = 0.091 µM) was pre-incubated with DAR (250
nM) for 0 – 80 min at 37 °C in the presence or absence of all-trans-retinaldehyde (250 μM)
before the enzyme reaction was initiated by addition of all-trans-retinaldehyde (250 μM) and
+
NAD (Figure 4D). RALDH2 activity was significantly decreased at 5 min (39.18 ± 6.10%
activity remaining), 10 min (30.43 ± 3.09% activity remaining), and 20 min (27.80 ± 2.87%
activity remaining) of pre-incubation, with maximal inhibition observed between 40 (16.31 ±
3
.98% activity remaining) and 80 min (16.50 ± 4.21% activity remaining) (***p < 0.001, one-
way ANOVA with Bonferroni’s multiple comparison test). Pre-incubation of RALDH2 with
vehicle alone (5 – 80 min at 37°C) had no significant effect on RALDH2 activity. No protection
of RALDH2 activity was observed when all-trans-retinaldehyde was included in the pre-
incubation period (250 nM DAR + 250 μM RAL). However, pre-incubation of RALDH2 with
+
DAR and NAD (250 µM) resulted in a recovery of 52 – 65% of RALDH2 activity as compared
#
#
###
with pre-incubation with DAR and RAL or DAR alone ( p<0.01, p<0.001, Student’s t-test).
1
1

Inhibitors that exhibit time dependence demonstrate a shift in IC values depending on the time
5
0
of pre-incubation which allows for the determination of the rate of inactivation (kinact) and
8
2-85
inhibition constant (KI).
Therefore, IC shift assays were performed and the kobs determined
50
from the slopes of lines calculated from the log of RALDH2 activity (log % Activity Remaining)
following 10 – 40 min of pre-incubation with DAR at concentrations of 0 – 500 nM. From these
slopes, the rate of inactivation (k ) was determined to be .000028 ± 0.00086 – 0.012 ± 0.009
obs
log % activity remaining/min for 0 – 500 nM DAR (Figure 4E). From the kobs values calculated
-1
in Figure 4E, the kinact and K for RALDH2 by DAR were calculated to be 0.0159 min and 128
I
-1
-1
nM, respectively with a time dependence ratio (kinact/K ) of 124,000 M min (Figure 4F). The
I
time dependence ratio calculated for DAR inactivation of RALDH2 is within the range reported
-1
-1
-1
-1 48
for the covalent inhibition of RALDH2 by DEAB (≈93,000 M min – 140,00,000 M min ).
RALDH1 and RALDH3 also exhibited a decrease in K and V with increasing concentrations
m
max
of DAR and time dependent inhibition, suggesting that DAR is an irreversible inhibitor for
chick RALDH1 and RALDH3 as well (Figure S1, Table S1). Interestingly, pre-incubation of
RALDH3 with retinaldehyde and DAR resulted in a significant increase in RALDH3
inactivation as compared with pre-incubation with DAR alone. These results suggest that the
irreversible binding of DAR to RALDH3 is somehow promoted by the presence of retinaldehyde.
At present, it is unclear as to the molecular basis for this interaction.
[insert figure 4 here]
2
.3. EX VIVO EFFECTS OF COMPOUND 2
1
2

(Dox)
RALDH2-eGFP, a HEK-293 cell line expressing doxycycline (DOX)-inducible
chicken RALDH2 was employed to assess the effect of DAR on intracellular RALDH2 activity
(Dox)
(Figure 5). Treatment of the
RALDH2-eGFP cells with DOX (5 μg/mL; 24 hrs) resulted in
(Dox)
the induction of RALDH2-eGFP expression as compared to non-induced
RALDH2-eGFP
(Dox)
cells (Figure 5A). In order to determine the toxicity of DAR on the
RALDH2-eGFP cells,
cell viability assays measuring DNA content and ATP production were performed (Figure 5B).
Following treatment with DAR (0 – 50 μM) for 24 hrs, toxicity curves were determined with IC50
values of 2.94 ± 1.58 μM and 3.09 ± 0.48 μM, for DNA and ATP production, respectively.
Treatment of DOX-induced RALDH2-eGFP expressing cells with DAR (0 – 2 μM for 24 hrs)
resulted in a significant inhibition of ATRA synthesis at 0.05 μM (↓42%, p<0.01), 0.50 μM
(
↓47%, p<0.001), 1 μM (↓92%, p<0.001), and 2 μM DAR (↓93%, p<0.001) compared to the
vehicle treated control (one-way ANOVA with Bonferroni’s test for multiple comparisons)
Figure 5C). No ATRA synthesis was detected in cells not induced with DOX (-DOX).
(
Normalization of results in Figure 5C to RALDH2 protein expression in each sample (relative
density quantified from western blot, inset) indicated a dose-dependent decrease in ATRA
synthesis with an IC value for the normalized inhibition curve calculated to be 187.20 ± 40.80
50
nM (Figure 5D).
[insert figure 5 here]
3
7,38,86
We and others
have previously shown that RALDH2 protein expression and
RALDH enzymatic activity are detectible in the choroids of chick eyes, with negligible amounts
in the retina/RPE and sclera. Moreover, ATRA synthesis is increased in the chick choroid during
recovery from form-deprivation induced myopia, which results from an increase in RALDH2
1
3

protein expression. Western blot analyses of choroidal lysates indicates that RALDH2 protein
↑125%
over contralateral control eyes). Increased RALDH2 protein expression in recovering choroids is
responsible for an 136% increase in RALDH enzymatic activity in choroidal lysates in
(ꢀ
ꢀ
recovering eyes. We therefore employed this animal model to examine the effect of DAR on
RALDH2 activity in choroids of recovering eyes and contralateral control eyes (Figure 6).
Choroidal cytosol fractions from control and 4 day recovering chick eyes were isolated and pre-
incubated with DAR (0 – 6 μM) for 20 min at 37°C prior to initiating the enzyme reaction by
+
addition of NAD (4 mM) and all-trans-retinaldehyde (25 μM), and RALDH2 activity was
measured by HPLC quantification of ATRA (Figure 6A). RALDH2 activity was significantly
inhibited in cytosol fractions from control and recovering choroids with 0.10 – 6 μM DAR (92 –
9
9% inhibition in control lysates and 70 – 96% inhibition in recovering lysates compared to
vehicle treated choroids, respectively (p < 0.001, one-way ANOVA with Bonferroni’s test for
multiple comparisons). Non-linear regression analyses of ATRA synthesis by choroid cytosol
fractions from recovering eyes following treatment with DAR indicated that DAR inhibited
RALDH2 activity in recovering choroid cytosol fractions with an IC of 53.60 nM (Figure 6B).
5
0
RALDH2 activity was also assessed, ex vivo, in choroidal lysates following overnight incubation
of living, intact choroids with DAR (0 – 5 µM) (Figure 6C, D). Choroids were isolated from 4
day recovering eyes or contralateral control eyes and incubated with DAR (0 – 5 μM) in organ
culture for 24 hrs, after which time choroid cytosol fractions were prepared and RALDH activity
measured. In contrast to results in which DAR was directly applied to choroid cytosol fractions
(in Figure 6A), DAR had no significant effect on ATRA synthesis in living, intact choroids from
control eyes. ATRA synthesis was maximally inhibited in recovering choroids by 1 μM (42%
inhibition), compared to vehicle treated choroids (p < 0.05, one-way ANOVA with Bonferroni’s
1
4

test for multiple comparisons). The EC50 (relative IC50) for this 42% inhibition of ATRA by
DAR was determined to be 119.70 nM when applied to living intact tissue, ex vivo (Figure 6D).
For comparison, living, intact choroids from control and 4 day recovering eyes were incubated
6
7,68
with WIN 18,446, a potent and irreversible ALDH1a inhibitor
(Figure 6E). Similar to
results with DAR, WIN had no significant effect on ATRA synthesis in living, intact choroids
from control eyes. However, ATRA synthesis was significantly inhibited in recovering choroids
by 0.1 μM WIN (40% inhibition), 1 µM WIN (51% inhibition) and 10 µM WIN (76% inhibition)
compared to vehicle treated choroids (p < 0.05, 0.01 and 0.001, respectively; one-way ANOVA
with Bonferroni’s test for multiple comparisons). Non-linear regression analyses of ATRA
synthesis by recovering choroids following treatment with WIN (Figure 6F) indicated that WIN
inhibits RALDH2 activity in recovering choroid lysates with an EC of 44.47 nM, when applied
5
0
to living intact tissue, ex vivo.
[insert figure 6 here]
2
.4. MOLECULAR MODELING.
We performed a series of molecular docking simulations using a homology model of
chicken RALDH2 constructed from the PDB crystal structure of sheep RALDH1 (PDB:
4
5
5
ABM) in order to investigate possible binding interactions of DAR. Based on the proposed
7
5
mechanism of inhibition by WIN 18446 of human RALDH2 and the known mechanism of
3
aldehyde oxidation by the ALDH enzymes, we predicted that an interaction would occur
1
5

between either the thiolate group of the catalytic cysteine of RALDH2 and either the carbonyl
carbon or carbon 1 of DAR (see Figure 1) resulting in an irreversible inhibition of the enzyme.
Binding poses were visualized with PyMOL to determine the most likely binding interaction by
examining the Van der Waals radii and any overlap that occurred between DAR and residues of
the RALDH2 active site (Figure 7). Three dimensional modeling of DAR in the binding pocket
of RALDH2 (Figure 7A) predicts the cyclic end group of DAR to be located at the entrance of
the substrate binding cavity, and carbon 1 of DAR with an attached chloride atom located deep
in the binding cavity in association with Cys302 of RALDH2. Moreover, the most probable
interaction occurs between the γ-sulfur of that catalytic cysteine and carbon 1 of DAR (cyan,
stick representation), resulting in the carbonyl carbon and chloromethyl end-group of DAR
being located deep in the substrate binding pocket, possibly participating in stabilizing hydrogen
bonding interactions with the residues of NAD, Asn169, and Cys302 (Figure 7B,C).
Interactions between the γ-sulfur of the catalytic cysteine of RALDH2 and the carbonyl carbon
of DAR were ruled out as modeling predicted insufficient space in the substrate binding pocket
to accommodate the additional methylene group of DAR. We predict that, similar to what has
75
been observed with WIN 18446, upon nucleophilic attack by the catalytic cysteine, a chloride
atom is displaced from DAR, resulting in an irreversible interaction (Figure 7C). Modeling of
+
87
DAR in the binding pocket of RALDH2 using the hydride transfer configuration of NAD
+
indicated significant atomic collision between the nicotinamide ring of NAD with the
dichloromethyl end group of compound 2, suggesting steric inhibition of DAR binding in the
+
active site of RALDH2 when NAD is present (data not shown). This model is substantiated by
results presented in Figure 4D, which demonstrated some protection of DAR inhibition by pre-
+
incubation with NAD .
1
6

[insert figure 7 here]
3
. CONCLUSIONS
Aldehyde dehydrogenases are essential for many biological processes and disease states.
In particular, the RALDH members of the ALDH family have been linked to a variety of
3
9, 40, 42
32, 33
37, 38
diseases such as cancer,
obesity,
and myopia
due to their importance in modulating
endogenous concentrations of ATRA. Of the three RALDH isoforms, RALDH2, and to a lesser
7
4, 79, 88
extent, RALDH1 are responsible for the majority of ATRA synthesis in postnatal tissues.
Therefore, the identification of inhibitors selective for the RALDH isozymes would enable
investigations on the roles of these enzymes in normal and disease processes, as well as provide
new therapeutic options for patients with the aforementioned diseases. By using an intelligent
drug design approach based on enzyme structure and substrate affinity, we report here the
development of a novel compound, DAR, that is effective at inhibiting the RALDH isozymes in
the nanomolar range, with no inhibition observed for ALDH2.
DAR was most effective at inhibiting chick RALDH2, followed by chick RALDH3 and
chick RALDH1, with IC ’s of 55 ± 10.71 nM, 161 ± 16.57 nM, and 435 ± 99.70 nM,
50
respectively. Additionally, DAR inhibited human RALDH2 (IC = 191.32 ± 36.77 nM). This is
5
0
not surprising, considering the full length amino acid sequences of chick and human RALDH2
are 95.8% identical (99.2% similar) and are 100% identical among residues lining the active site.
Future studies comparing DAR on all human RALDH isoforms will provide necessary
information on isoform specificity of DAR for application to specific clinical conditions. Based
on the three dimensional structures of our chick RALDH models, RALDH1 contains a
methionine (Met120) residue that partially projects into the active site tunnel, potentially limiting
1
7

access of RALDH1 to DAR. Interestingly, this methionine is not found in RALDH2 or
RALDH3 (valine and isoleucine occupy this position, respectively), indicating that the access
tunnels of RALDH2 and RALDH3 are unobstructed and can readily accommodate DAR,
resulting in the observed lower IC values for DAR on RALDH2 and RALDH3, as compared
5
0
with RALDH1.
Several lines of evidence presented in this report indicate that DAR inhibits RALDH2 in
an irreversible and time-dependent manner, similar to the mechanism described for WIN 18446.
6
7, 68
75
Based on the proposed mechanism of inhibition of WIN 18446 on human RALDH2, we
predict that upon binding of DAR, nucleophilic attack on carbon 1 of DAR (see Figure 1) by the
thiolate group of the catalytic cysteine may result in the displacement of one of the chloride
atoms, resulting in the formation of a covalent, tetrahedral 2-chloro-2-(cysteinyl)acetaldehyde.
However, the chemical electrophilic reactivity of the terminal carbon atom may be different
between WIN 18446, which contains amide groups adjacent to the carbonyl carbons and DAR,
which contains a conjugated double bond system. Mass spectrometry and protein crystallization
experiments are underway to confirm the proposed mechanism of RALDH inhibition by DAR.
DAR was also evaluated in cell-based studies using a DOX-inducible chicken RALDH2
(Dox)
cell line ( RALDH2-eGFP), in choroid tissue homogenates, and in ex vivo studies with
isolated chick choroids. In all of these systems, DAR was effective at inhibiting ATRA
synthesis with IC values similar to that calculated in vitro for recombinant RALDH2 and well
50
below concentrations determined to be toxic, both in vitro and ex vivo. Interestingly, when
evaluating DAR on living intact choroids ex vivo, DAR was only effective on choroids isolated
from chick eyes during recovery from induced myopia (“recovering choroids”). We have
previously established that RALDH2 expression and activity are increased significantly in
1
8

37
recovering choroids and that RALDH2 is responsible for elevated ATRA concentrations in
3
8, 86
recovering choroids.
Moreover, when placed in vitro, only recovering choroids are capable
37
of de novo ATRA synthesis. The EC value for inhibition of ATRA formation in living
5
0
recovering choroids by compound 2 was estimated to be 119.7 nM, but DAR only inhibited a
maximum of 51% of ATRA formation in recovering choroids (with maximum inhibition at 1
µM). We speculate that DAR may be partially metabolized in living choroids, similar to the
8
9
metabolism of ATRA by the cytochrome p450 (CYP) 26 enzyme. This idea is strengthened by
our observation that DAR is a more effective ATRA synthesis inhibitor in choroid cytosol
fractions (in which microsome-associated enzymes were removed by centrifugation at 100,000 x
g) of both control and recovering eyes (IC = 20.7 nM and 53.6 nM, respectively). Moreover,
5
0
WIN 18,446 (WIN), a non-specific ALDH inhibitor of the bisdichloroacetyldiamine class of
compounds, is significantly more effective at inhibiting ATRA synthesis by recovering choroids
ex vivo (76% inhibition at 10 µM WIN; EC = 44.47 nM) as compared with DAR. DAR was
5
0
(Dox)
more effective at inhibiting RALDH2 activity in the HEK-293 cell line,
RALDH2-eGFP, as
compared with the level of RALDH2 inhibition in lysates of recovering choroids following
incubation of living choroids with DAR, ex vivo. It is possible that one or more cell type in the
(Dox)
chick choroid expresses higher levels of CYP 26 as compared with
RALDH2-eGFP, It is also
possible that limitations in cellular uptake and chemical instability of DAR may contribute to its
less effective action on living intact choroids. Therefore, these results suggest that for effective
inhibition of ATRA synthesis in vivo, larger concentrations of DAR may be needed to maximize
the inhibitory effect of DAR on RALDH2 and/or necessitate modifications to the structure of
DAR to limit its metabolism by CYP degradation, improve compound stability and increase
bioavailability.
1
9

4
. EXPERIMENTAL SECTION
4
.1. GENERAL METHODS FOR CHEMISTRY.
All reactions were performed in oven-dried glassware under a positive pressure of nitrogen
unless noted otherwise. Flash column chromatography was performed as described by Still et
9
0
al. employing E. Merck silica gel 60 (230 – 400 mesh ASTM). TLC analyses and preparative
TLC (pTLC) purification was performed on 250µm Silica Gel 60 F254 plates purchased from
EM Science (Hatfield, PA) and Fluka Analytical (St. Louis, MO). All solvents and chemicals
were used as purchased without further purification. Methylene chloride (CH Cl ) was prepared
2
2
by distillation with CaH . Infrared spectra were recorded on a Shimadzu IRAffinity-1 instrument
2
(Shimadzu; Kyoto, Japan), and IR spectra peaks are reported in terms of frequency of absorption
-1
1
13
(
cm ). H and CNMR spectra were recorded on VNMRS 400 and VNMRS 500 MHz-NMR
Spectrometers (Varian, Inc.; Paolo Alto, CA). Chemical shifts for proton and carbon resonances
are reported in ppm (δ) relative to the residual proton or the specified carbon in chloroform (δ
7
.27, proton; 77.23, carbon). High-resolution mass spectrometry (HRMS) analysis was
performed using Agilent 6538 high-mass-resolution QTOF mass spectrometer (Santa Clara, CA).
HPLC purification was performed on Shimadzu LCMS 2020 system [LC-20AP (pump), SPD-
M20A (diode array detector), LCMS-2020 (mass spectrometer)]. Semi-preparative HPLC
purification was performed using Phenomenex (Torrance, CA) Luna C-18(2) column, 5 µm
particle size (250 mm x 10mm), supported by Phenomenex Security Guard cartridge kit C18 (4.0
mm x 3.0 mm); and HPLC-grade solvents. Purity was determined by reverse-phase HPLC
(Phenomenex Luna 5µ C18(2) 100Ǻ, 250 mm x 10 mm). All compounds tested present a purity
>95%. Method: acetonitrile/water with 0.1% formic acid gradient (flow rate: 2.5 mL/min). The
2
0

HPLC run started with 96% acetonitrile for 3 minutes, followed by a gradient increase in
acetonitrile from 96-98% over 9 minutes, followed 98% acetonitrile for 8 minutes.
4
.2. (3E,5E,7E,9E)-1,1-DICHLORO-4,8-DIMETHYL-10-(2,6,6-TRIMETHYLCYCLOHEX-1-
EN-1-YL)DECA-3,5,7,9-TETRAEN-2-ONE (DAR): Note: DAR can photoisomerize under
ambient light conditions. All steps were performed in a darkened fume hood, and DAR was
wrapped in aluminum foil at all steps of purification to protect it from light. The stock solution
of DAR dissolved in DMSO was also protected from light at all times.
Ethyl retinoate (69.5 mg, 0.211 mmol) was dissolved in dry ethyl ether (0.9 mL) under
nitrogen, and dichloromethane (19.0 µL, 0.296 mmol) was added via gas-tight syringe. The
resulting dark yellow solution was cooled to -78˚C. Lithium diisopropylamide (LDA) solution
[2.0 M in tetrahydrofuran (THF), 170 µL, 0.338 mmol] was then added dropwise slowly over 5
min. Upon addition of LDA, the color of the reaction mixture turned dark red/brown. The
progress of the reaction was monitored by TLC (10% EtOAc/hexanes, CAM stain) every 2 min
for 6 minutes. The reaction mixture was then stirred at -78˚C for an additional 10 min, until no
further reaction progress was observed with TLC. The reaction mixture was quenched by
addition of 400 µL of 6N HCl. The mixture was then warmed to room temperature (RT), diluted
with ethyl ether (30 mL), washed with 1N HCl (15 mL x 2), followed by washes with distilled
water (15 mL x 2). The combined aqueous phase was back extracted with ethyl ether (5 mL x
2
). The combined organic phase was then washed with brine (20 mL) and dried over Na SO .
2
4
The solvent was removed under reduced pressure to afford the crude product as a dark brown oil
70 mg).
(
The reaction did not go to completion, and produced multiple products based on TLC
analysis. The crude mixture was first separated using preparative silica gel TLC (3 plates) and
2
1

eluted with 4.5% EtOAc/hexane. Band 1 (R = 0.54, 12.8 mg, desired product, DAR); band 2 (R
f
f
=
0.48, 14.9 mg, recovered ethyl retinoate); band 3 (R = 0.37, 7 mg, unassigned product); band 4
f
(
R = 0.22, 2.8 mg, mixture of compounds). The desired product, DAR, (Figure S2-S7) obtained
f
from pTLC fraction 1 was purified with reverse-phase HPLC using a Phenomenex C18(2) Luna
semiprep column with a 96 – 98% acetonitrile/0.1% aqueous formic acid solvent gradient. 2D-
1
1
1
13
NMR methods ( H- H COSY and H- C HMBC) were used to assign NMR signals to the
proposed structure. Pure desired product (DAR) was obtained as a yellow oil (7.7 mg, 10% yield,
1
1
2.6% borsm). DAR: R = 0.54 (4.5% EtOAc/hexanes); H NMR (500 MHz, CDCl ): δ = 7.21
f
3
(
d, J = 15.0, 11.5 Hz, 1H, H6), 6.49 (s, 1H, H3), 6.42 (d, J = 15.1 Hz, 1H, H5), 6.37 (d, J = 18.0
Hz, 1H, H7), 6.20 (d, J = 11.5 Hz, 1H, H9), 6.17 (d, J = 15.9 Hz, 1H, H10), 5.84 (s, 1H, H1),
.44 (s, 3H, H17), 2.05 (s, 3H, H18), 2.04 (m, 2H, H13), 1.73 (s, 3H, H19), 1.68 - 1.59 (m, 2H,
2
1
3
H14), 1.52-1.46 (m, 2H, H15), 1.05 (s, 6H, H20 and H21). C NMR (126 MHz, CDCl ): δ =
3
1
1
3
86.46 (C2), 159.02 (C4), 141.93 (C8), 137.63 (C11), 137.05 (C10), 134.81 (C5), 134.38 (C6),
30.63 (C12), 130.03 (C7), 129.40 (C9), 118.02 (C3), 71.02 (C1), 39.60 (C15), 34.28 (C16),
3.17 (C13), 28.97 (C20, C21), 21.77 (C19), 19.18 (C14), 14.96 (C17), 13.05 (C18). IR (NaCl,
-1
cm ): 3440 (C –H), 2920 (C –H), 1680 (C=O), 1558 (C=C), 785 (C–Cl). HRMS (ESI): m/z
sp3
sp2
calculated for C H Cl O + H+ [M + H+]: 367.1595, found: 367.1596, Δ = 0.27 ppm. Purity
2
1
28
2
was determined as >95% by HPLC. R : 16.8 min.
t
4
.3. BIOLOGICAL TESTING OF DAR
DAR is prone to photoisomerization, and therefore, was protected from ambient light at
all times. DAR was dissolved in dimethylsulfoxide (DMSO) to produce a 10 mM solution for
biological testing. These stock solutions were protected from light and stored frozen at -20ºC.
2
2

Multiple small aliquots of DAR (10 mM) were prepared and used for biological testing to
prevent refreezing of sample as DAR will degrade upon frequent freeze/thaw cycles and
exposure to ambient light.
are indicative of degradation and loss of function. ꢁherefore we have determined that this
compound is stable, if stored in the darꢂ and at -80 for at least three months. Fresh aliquots of
DAR uniquely absorbs at 410 nm, and shifts in the spectral profile
ꢃ
DAR were used for each assay and DAR was stable under assay conditions (darꢂ, 25 ꢃ C and 3ꢄ ꢃ
C) for at least three days.
4
.4. ALDH1A/ALDH2 EXPRESSION AND PURIFICATION
Chicken recombinant RALDH1 (ALDH1A1), RALDH2 (ALDH1A2), RALDH3
3
8
(
ALDH1A3), were produced and purified, as previously described. Human recombinant
ALDH2 (hALDH2) and human recombinant RALDH2 (hRALDH2) were prepared from
plasmids generously provided by Dr. Jisun Paik (University of Washington, U.S.A.), and protein
was produced as described for chicken recombinant proteins except that kanamycin (250 µg/ml)
was used as the bacterial selection agent. Following purification on cobalt-agarose columns,
RALDH proteins were dialyzed in 20 mM triethanolamine-HCl, pH 7.4 containing 0.1 mM
EDTA (buffer degassed prior to use), while hALDH2 was dialyzed in 20 mM Hepes, pH 8.5
containing 150 mM KCl and 1 mM EDTA. After dialysis, 500 mM tris(2-carboxyethyl)
phosphine (TCEP) (Thermo Fisher Scientific; Waltham, MA) was added to the protein solutions
67
to a final concentration of 1 mM. Purified enzymes were stored at 4 °C for up to 6 months, and
enzyme stability/activity was measured at least once a month. When stored in 1 mM TCEP,
recombinant RALDH enzymes demonstrated a ≤ 40% loss of activity after 6-month storage at
4
°C; therefore, enzyme assays were routinely carried out within 2 months following protein
purification.
2
3

4
.5. ENZYME ACTIVITY ASSAYS USING PROPIONALDEHYDE
Dehydrogenase activity of purified recombinant hALDH2 was assayed
spectrophotometrically by monitoring the formation of NADH at 340 nm using a microplate
reader (FLUOstar OPTIMA, BMG Labtech; Ortenberg, Germany). Recombinant hALDH2 (4.5
μM in dialysis buffer, see above) was added to 96 well plates with assay buffer [20 mM Hepes
pH 8.3, 150 mM KCl, 1 mM EDTA, 2 mM dithiothreitol (DTT), 5 mM propionaldehyde in
EtOH; prepared from a 2x stock] in a volume of 100 μL at 25°C. Reactions were initiated by
+
adding NAD to a final concentration of 5 mM (from a 20 mM stock in dH O), and the rate of
2
NADH synthesis was monitored at 340 nm for 5 min at 25°C. When present, ALDH inhibitors (1
–
100 µM in DMSO, final concentration) or vehicle (DMSO) were added to 96 well microplates
+
and pre-incubated (20 min – 1 hr) with hALDH2 and assay buffer prior to addition of NAD and
monitoring of NADH synthesis at 340 nm.
4
.6. ENZYME ACTIVITY ASSAYS USING RETINALDEHYDE
Due to overlap in the absorption spectrum of retinaldehyde and NADH, a fluorescence-
based NADH assay (Amplite™ Fluorimetric NADH Assay Kit, AAT Bioquest, Inc; Sunnyvale,
CA) was employed to measure dehydrogenase activity of purified recombinant RALDH
isozymes with the natural substrate, retinaldehyde. Recombinant protein was diluted in dialysis
buffer (see above) containing 2 mM DTT to a volume of 40 µL and added to amber microfuge
+
tubes containing 50 µL of 2x assay buffer [5% DMSO, 8 mM NAD , 64 mM tetrasodium
pyrophosphate pH 8.2, 0.2 mM pyrazole, 10 mM reduced glutathione (GSH), 1.9 mM EDTA] at
2
5°C. Retinaldehyde was used directly from sealed ampules (Sigma, ≥98% purity, 25 mg/vial),
stored at -20° and discarded after 1 week. Retinaldehyde was initially solubilized in ethanol and
its concentration determined by absorption spectroscopy (λmax = 383 andε= 42,880) of serial
2
4

9
1
dilutions. The retinaldehyde stock solution was then diluted to 10 x concentrations (0.155 mM
-
2.50 mM) in ethanol. Under dim red light, reactions were initiated by addition of retinaldehyde
in EtOH (10 uL; final retinaldehyde concentration = 15.5 - 250 μM; final reaction volume = 100
µl), and reactions were incubated in a water bath for 30 min at 37°C. Under these conditions,
retinaldehyde remained in solution in synthesis buffer at all concentrations. Reactions were
stopped by immersion in ice water. For NADH quantification, a 50 µl aliquot of each reaction
was added to wells of solid black 96 well microplates (Greiner Bio One; Kremsmünster, Austria)
followed by 50 µl of the NADH reaction mixture (Amplite™ Fluorimetric NADH Assay Kit,
according to manufacturer’s directions). Total NADH was quantified by fluorescence intensity
at Ex/Em = 540/590 nm and compared with a standard curve (0.1 – 30 µM NADH). While an
+
excess of NAD was added to the reactions, the amount of NADH generated in the 30 min was
within the range of the standard curve. For inhibition assays, recombinant protein was pre-
incubated with DAR at a final concentration of 0 – 10 μM in DMSO for 0 – 80 minutes at 37°C
prior to addition of assay buffer and retinaldehyde (DMSO final concentration after addition of
substrate and synthesis buffer ≤ 3.5%). The calculations for K and kinact were performed as
I
8
4, 85
previously described.
4
.7. CELL CULTURE AND DAR TREATMENT
(Dox)
A stably transfected HEK-293 cell line,
RALDH2-eGFP, that exhibits doxycycline-
inducible expression of chicken RALDH2-eGFP was employed to assess the effect of DAR on
(Dox)
intracellular RALDH2 activity. The
RALDH2-eGFP cell line was generated via lentiviral
infection of HEK 293 cells as follows: The lentiviral transfer vector, #143, was generated by
ligation of the chicken RALDH2 coding sequence (GeneBank Accession # NM_204995)
together with the ribosomal entry site (IRES) and eGFP (obtained from pTet-IRES-EGFP;
2
5

9
2
plasmid #64238 Addgene, Cambridge, MA) into the open reading frame of pInducer 20
(obtained as a generous gift from Dr. Thomas Westbrook, Baylor College of Medicine, Houston
TX). This construct encoded for constitutive expression of the transactivator (rtTA3) and
neomycin resistance (Neo) via the Ubc promoter as well as inducible expression of RALDH2-
IRES-eGFP by a tetracycline responsive element (TRE2) in the presence of doxycycline. Third
generation lentiviral particles were generated by the transient co-transfection of HEK 293T with
the lentiviral transfer vector plasmid (#143; 8 µg/10 cm plate), packaging plasmids (pRSV-Rev
and pMDL/pRRE; 2µg each/ 10 cm plate) and envelope plasmid DNA (pMD2.G; 2 μg/10 cm
plate) together with an equal mass of PEI (14 µg/10 cm plate, from a 2 ug/ul stock) in Opti-
MEM (Gibco® by Life Technologies, Carlsbad, CA) at a final volume of 1 ml DNA solution per
1
0 cm plate. DNA/PEI mixtures were incubated at room temperature for 15 min added to
dropwise to plated 293T cells (1ml per 10 cm plate) and incubated at 37°C in a 5%
CO humidified atmosphere. The culture supernatants containing lentiviral particles were
2
collected at 48 – 72 hrs post-transfection, concentrated by ultracentrifugation at 20,000 rpm
(Beckman rotor SW-28) for 2.5 hrs at 4 °C and stored at −80°C. A stable cell line expressing
inducible chicken RALDH2 and eGFP was subsequently generated by lentiviral transduction of
HEK 293 cells in the presence of polybrene (8 μg/mL). Transfected cells were monitored by
eGFP fluorescence and sorted via flow cytometry using a BD Biosystems LSRII analyzer.
RALDH2-eGFP positive cells were cultured in Dulbecco’s modified Eagle medium
(DMEM)/high glucose (Thermo Fisher Scientific, Waltham, MA, USA) supplemented with 10%
fetal bovine serum (FBS; GE Healthcare Bio-Sciences, Pittsburgh, PA, USA) containing
neomycin sulfate (1 µg/ml). The cells were maintained at 37°C and 5% CO in a humidified
2
2
6

incubator (VWR International, LLC., Atlanta, GA, USA) and were harvested at 80–90%
confluence.
(Dox)
For experiments testing DAR,
a density of 150,000 cells/well in culture medium [DMEM containing 10% fetal bovine serum
FBS) and doxycycline (5 µg/ml)] and incubated at 37°C and 5% CO for 25 hrs. Culture
RALDH2-eGFP cells were seeded in 6- well plates at
(
2
medium was replaced with low serum media (DMEM containing 0.1% FBS) containing 0.05 – 2
µM DAR in DMSO (< 0.004%) or vehicle, and cells were incubated for 8 hrs at 37°C and 5%
CO . Following incubation, media was removed, and cell layers were collected [1.0 ml + 0.5 ml
2
homogenization buffer (20 mM triethanolamine-HCl pH 7.4, 1 mM DTT, 0.1 mM EDTA)] with
the aid of a rubber policeman. Cells were lysed using a VirTis rotor-stator homogenizer (SP
Industries; Gardiner, NY). Homogenates were then transferred to thick-walled microfuge tubes
(
polyallomer tubes; Beckman Coulter; Brea, CA) and ultracentrifuged (100,000 x g for 1 hr;
Optimum MAX Ultracentrifuge, Beckman Coulter) at 4°C to isolate the cytosol fraction
supernatant) which was stored at -20°C until use. Relative RALDH2 protein expression was
(
38
determined by western blot, as previously described. RALDH2 protein expression using the
(Dox)
RALDH2-eGFP 293 HEK cell line ranged from 25% – 37% following incubation with 0.05
–
2 µM DAR, as determined from three independent experiments, each performed in triplicate.
RALDH activity was determined by measuring the production of ATRA in vitro, as
38
described previously. All procedures with retinaldehyde or ATRA were performed under dim
red light. 200 μL of synthesis buffer [2.5% DMSO, 4 mM NAD, 32 mM tetrasodium
pyrophosphate pH 8.2, 0.1 mM pyrazole, 5 mM reduced glutathione (GSH), 1 mM EDTA;
prepared from a 2x stock] was added to 200 μL of cytosol from homogenized cells. The reaction
was initiated by the addition of 50 μL of 250 μM all-trans-retinaldehyde (in synthesis buffer) to
2
7

the reaction mixture (25 μM final concentration). The reactions were vigorously mixed and
placed in a water bath at 37°C for 30 min, after which time, the reactions were stopped by
immersion in ice water and addition of 3/2 volumes (675 μL) of methanol. Samples were stored
at -80°C until time of use.
91
ATRA was extracted for quantification by reverse phase HPLC, as previously described
with the following modifications: to 500 μL of sample, 60 μL of acetonitrile and 17 μL of 4M
HCl were added to clear, microcentrifuge tubes and vortex mixed for ~ 15 sec. Hexane (600 μL)
was added to each tube and vortex mixed for ~ 10 sec. Tubes were then centrifuged at RT for 3
min at 1000g to facilitate phase separation. The organic (top) phase was removed (400 μL), with
great care taken to not disturb the bottom aqueous layer, and transferred to a new, clear
microcentrifuge tube (SafeSeal; Sorenson BioScience Inc, Salt Lake City, UT). The organic
phase was then evaporated under a gentle nitrogen stream at RT. Samples were then
resuspended in 100 μL of acetonitrile and transferred to plastic HPLC vials (9 mm S/T vial with
septa) (Supelco, Inc; Bellefonte, PA). All steps were performed in dim red light. ATRA was
9
0
quantified as previously described with the following modifications: a Shimadzu LCMS 2020
(Shimadzu; Kyoto, Japan) with a Cortecs C18 column [2.7 μm particle size with VanGuard
Cartridge (Waters; Milford, MA)] was used to achieve ATRA separation. The column
compartment was maintained at 25°C, and the autosampler was maintained at 10°C. The
following solvents were used: A, H O with 0.1% formic acid; B, acetonitrile. The column was
2
equilibrated in 70% acetonitrile prior to sample loading. 25 μL of sample was injected into the
column, and separation was achieved at 400 μL/min with the following gradient: 0 – 3 min, hold
at 70% B; 3 – 15 min, 70% B to 95% B; 15 – 20 min, hold at 95% B; 20 – 21 min, 95% B to
7
0% B; 21 – 37 min, re-equilibrate at 70% B. Elution of ATRA was monitored using a
photodiode array detector at 350 nm, followed by MS detection. Retention time of ATRA was
2
8

2
0.6 min with an m/z of 300.25. ATRA was quantified from a standard curve (Figure S8)
generated using known amounts of ATRA (0 – 1500 pmol) eluted with the above gradient.
For cell viability assays, aliquots of the cell homogenates (prior to ultracentrifugation)
were digested with proteinase K (protease type XXVIII, Sigma; in 10 mM EDTA, 0.1 M sodium
phosphate pH 6.5; 0.05% w/v) at 60°C overnight (O/N). DNA was measured directly from
proteinase K digests using Pico-green (Invitrogen; Carlsbad, CA) and a mini-fluorometer (TBS-
3
80; Turner Biosystems; Sunnyvale, CA), according to manufacturer’s protocols. Additionally,
cell viability was determined by quantification of ATP in separate cell cultures. For ATP
(Dox)
quantification,
RALDH2-eGFP 293 cells were plated at a density of 10,000 cells/well solid
black, flat-bottomed 96 well microplates (Greiner Bio One; Kremsmünster, Austria) in culture
medium (DMEM containing 10% FBS) and incubated at 37°C and 5% CO for 24 hrs. Culture
2
medium was replaced with low serum media (DMEM containing 0.1% FBS) containing 0 – 50
µM DAR in DMSO (< 0.5% DMSO) and incubated for 24 hrs at 37°C and 5% CO . ATP was
2
quantified using the CelTiter-Glo Luminescent Cell Viability Assay (Promega; Madion, WI)
according to manufacturer’s instructions. Luminescence was measured using a FLUOstar
OPTIMA plate reader (BMG Labtech; Cary, NC). Viability was calculated as a percentage of
vehicle-treated (0.5% DMSO v/v) control cells at each concentration of DAR.
4
.8 ANIMALS
White Leghorn male chicks (Gallus gallus) were obtained as 2 day old hatchlings from
Ideal Breeding Poultry Farms (Cameron, TX). Chicks were housed in temperature controlled
brooders with a 12-hour light/dark cycle and were given food and water ad libitum. Form
deprivation myopia (FDM) was induced in 3 day old chicks by applying translucent plastic
93
goggles to one eye, as described previously. The contralateral eyes (left eyes) of all chicks were
2
9