Question

The human pathway for metabolizing alcohol starts with the enzyme alcohol dehydrogenase, which catalyzes the conversion of ethanol (C2H5OH) to acetaldehyde (CH3CHO). This is followed by the aldehyde dehydrogenase 2 (ALDH2, the enzyme of interest in this problem), which catalyzes the conversion of acetaldehyde and HS-CoA to acetyl-CoA (CH3CO–S–CoA). The TCA cycle starts and oxidizes the acetyl-CoA to CO2. Draw two diagrams of this pathway—one for an individual without AFS and another for an individual with AFS. How ALDH2 deficiency combined with ethanol input into the bloodstream could culminate in the accumulation of acetaldehyde in the blood of the patient.

Answer 1

This is the explaintion that

The primary enzymes involved in alcohol metabolism are alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH). Both enzymes occur in several forms that are encoded by different genes; moreover, there are variants (i.e., alleles) of some of these genes that encode enzymes with different characteristics and which have different ethnic distributions. Which ADH or ALDH alleles a person carries influence his or her level of alcohol consumption and risk of alcoholism. Researchers to date primarily have studied coding variants in the ADH1B, ADH1C, and ALDH2 genes that are associated with altered kinetic properties of the resulting enzymes. For example, certain ADH1B and ADH1C alleles encode particularly active ADH enzymes, resulting in more rapid conversion of alcohol (i.e., ethanol) to acetaldehyde; these alleles have a protective effect on the risk of alcoholism. A variant of the ALDH2 gene encodes an essentially inactive ALDH enzyme, resulting in acetaldehyde accumulation and a protective effect. It is becoming clear that noncoding variants in both ADH and ALDH genes also may influence alcohol metabolism and, consequently, alcoholism risk; the specific nature and effects of these variants still need further study.....

Alcohol and other drug (AOD) use (AODU), abuse and dependence; alcoholism; genetics and heredity; genetic theory of AODU; ethnic group; protective factors; ethanol metabolism; liver; alcohol dehydrogenase (ADH); aldehyde dehydrogenase (ALDH); risk factors; protective factors; alcohol flush reaction

The effects of ingested beverage alcohol (i.e., ethanol) on different organs, including the brain, depend on the ethanol concentration achieved and the duration of exposure. Both of these variables, in turn, are affected by the absorption of ethanol into the blood stream and tissues as well as by ethanol metabolism .The main site of ethanol metabolism is the liver, although some metabolism also occurs in other tissues and can cause local damage there. The main pathway of ethanol metabolism involves its conversion (i.e., oxidation) to acetaldehyde, a reaction that is mediated (i.e., catalyzed) by enzymes known as alcohol dehydrogenases (ADHs). In a second reaction catalyzed by aldehyde dehydrogenase (ALDH) enzymes, acetaldehyde is oxidized to acetate. Other enzymes, such as cytochrome P450 (e.g., CYP2E1), metabolize a small fraction of the ingested ethanol.There are multiple ADH and ALDH enzymes that are encoded by different genes .Some of these genes occur in several variants (i.e., alleles1), and the enzymes encoded by these alleles can differ in the rate at which they metabolize ethanol or acetaldehyde or in the levels at which they are produced. These variants have been shown to influence a person’s drinking levels and, consequently, the risk of developing alcohol abuse or dependence Studies have shown that people carrying certain ADH and ALDH alleles are at significantly reduced risk of becoming alcohol dependent. In fact, these associations are the strongest and most widely reproduced associations of any gene with the risk of alcoholism. As will be discussed later in this article, the alleles encoding the different ADH and ALDH variants are unevenly distributed among ethnic groups.

Methods

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Methods will be Wild type and ALDH2 deficient mice were fed (1–6%) in Lieber-DeCarli diet for 4 weeks. Gut permeability in vivo measured by plasma-to-luminal flux of FITC-inulin, tight junction and adherens junction integrity analyzed by confocal microscopy and liver injury was assessed by analysis of plasma transaminase activity, histopathology and liver triglyceride...

Result

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In Ethanol feeding elevated colonic mucosal acetaldehyde, which was significantly greater in ALDH2 deficient mice. ALDH2−/− mice showed a drastic reduction in the ethanol diet intake. Therefore, this study was continued only in wild type and ALDH2+/− mice. Ethanol feeding elevated mucosal inulin permeability in distal colon, but not in proximal colon, ileum or jejunum of wild type mice. In ALDH2+/− mice, ethanol-induced inulin permeability in distal colon was not only higher than that in wild type mice, but inulin permeability was also elevated in the proximal colon, ileum and jejunum. Greater inulin permeability in distal colon of ALDH2+/− mice was associated with a more severe redistribution of tight junction and adherens junction proteins from the intercellular junctions. In ALDH2+/− mice, but not in wild type mice, ethanol feeding caused a loss of junctional distribution of tight junction and adherens junction proteins in the ileum. Histopathology, plasma transaminases and liver triglyceride analyses showed that ethanol-induced liver damage was significantly greater in ALDH2+/− mice compared to wild type mic.

I will show you the tables

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Alcohol Dehydrogenase (ADH) Genes and Proteins

Official Gene Name*	Old Name†	Nonstandard Name‡	Sequence§	Protein	Class¶
ADH1A	ADH1	ADH1A	NM_000667	α	I
ADH1B	ADH2	ADH1B	NM_000668	β	I
ADH1C	ADH3	ADH1C	NM_000669	γ	I
ADH4	ADH4	ADH2	NM_000670	π	II
ADH5	ADH5	ADH3	NM_000671	χ	III
ADH6	ADH6	ADH5	NM_000672	ADH6	V
ADH7	ADH7	ADH4	NM_000673	σ	IV

Kinetic properties are given below;

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Kinetic Properties of Alcohol Dehydrogenase (ADH) Proteins

Official Gene Name*	Amino Acid Differences Between Alleles	Protein Name	Km(ethanol) mM	Turnover (min−1)
ADH1A		α	4.0	30
ADH1B*1	Arg48, Arg370	β1	0.05	4
ADH1B*2	His48, Arg370	β2	0.9	350
ADH1B*3	Arg48, Cys370	β3	40	300
ADH1C*1	Arg272, Ile350	γ1	1.0	90
ADH1C*2	Gln272, Val350	γ2	0.6	40
ADH1C*352Thr	Thr 352†	—	—	—
ADH4		π	30	20
ADH5		χ	>1,000	100
ADH6		ADH6	?	?
ADH7		σ	30	1800

ALDH2 expression in liver

Human ALDH2 mRNA levels in the liver were assessed by TaqMan RT-qPCR using FAM dye labeled hALDH2-specific primer–probe sets (Assay ID; Hs01007998_ml) and murine 18S-specific primer–probe sets (Life Technologies, Waltham, MA). ALDH2 protein was quantified by Western analysis with an antibody against the HA tag (Sigma-Aldrich, St. Louis, MO). Paraformaldehyde-fixed livers were embedded in paraffin and cut into 5 μm sections. Vector-derived hALDH2 expression in the liver was detected by an anti-HA antibody (Histowiz, Brooklyn, NY). After sectioning, anti-HA immunohistochemical staining, and counterstaining with hematoxylin, digital images of liver cross-sections were acquired using a 20 × objective. The enzymatic activity of ALDH2 in liver was analyzed using the colorimetric mitochondrial aldehyde dehydrogenase (ALDH2) activity assay kit (ab115348; Abcam, Cambridge, MA) according to the manufacturer's protocol using 200 μg of total protein from liver homogenate. C57Bl/6 mice administered PBS were used as controls.

Acetaldehyde

Acetaldehyde measurements were performed at the Proteomics Resource Center, Rockefeller University9,34 (n = 5 mice/group). Acetaldehyde levels were determined through derivatization with dinitrophenylhydrazine (DNPH) using butyraldehyde–DNPH (Accustandard, New Haven, CT) as an internal standard. Blood (120 μL) was deproteinized by addition of an equal volume of cold acetonitrile (ACN) and centrifuged for 30 min at 3,500 g at 4°C. Supernatant (48 μL) was mixed with 2 μL of 10 mM 13C-acetaldehyde. Then, 15 μL of 16 mM DNPH in ACN and 5 μL of 1 M citric acid (pH 4.0) were added. After incubation at 25°C for 30 min, the reaction was quenched with 60 μL of 0.1% formic acid in water, and samples were kept in −80°C until analysis. Before LC/MS, 49 μL of sample was spiked with 1 μL of butyraldehyde–DNPH standard for a final concentration of 20 pm/μL. Acetaldehyde–DNPH (Accustandard, New Haven, CT) was used as an external calibrant to verify retention time. Samples were analyzed on a Dionex U-3000 HPLC system coupled to a TSQ Vantage triple-quad mass spectrometer (Thermo Fisher Scientific, Waltham, MA) equipped with a heated electrospray ionization source (HESI). Chromatographic separation was performed using a Thermo Acclaim 120 C8 (2.1 × 150 mm, 3 μm particle size) column at a flow rate of 200 μL/min, using 0.1% LC-grade formic acid (Pierce, Thermo Fisher Scientific) in water as buffer A and 100% methanol (Optima, Fisher Scientific, Hampton, NH) as buffer B. The gradient was the following: 5% buffer B (0–2 min), increased to 60% buffer B (2–10 min), increased to 80% buffer B (10–20 min), increased to 100% buffer B (20–21 min), held at 100% buffer B (21–26 min), returned to 50% buffer B (26–26.1 min) and held at 50% buffer B (26.1–30 min) for re-equilibration of the column. The mass spectrometer was operated with the following parameters; negative ion polarity; spray voltage, 3500 V; ion transfer capillary temperature, 350°C; source temperature, 37°C; sheath gas, 35 (arbitrary units); auxiliary gas, 10 (arbitrary units); collision gas, argon. The S-Lens voltage (S-lens) and collision energy (CE) were manually optimized for each multiple reaction monitoring (MRM) transition. The MRM transitions monitored were: Acetaldehyde-DNPH, m/z 223→122 (CE = 21, S-Lens = 65), 223→163 (CE = 13, S-Lens = 65), 223→181 (CE = 18, S-Lens = 65); 13C-Acetaldehyde-DNPH, m/z 225→122 (CE = 21, S-Lens = 65), 225→163 (CE = 13, S-Lens = 65), 225→181 (CE = 18, S-Lens = 65); Butyraldehyde-DNPH, m/z 251→122 (CE = 27, S-Lens = 70), 251→152 (CE = 19, S-Lens = 70), 251→163 (CE = 13, S-Lens = 70). Quantitation was performed using a known amount of 13C-acetaldehyde standard in the sample and an external calibration curve of acetaldehyde–DNPH prepared in parallel. Rate constants of acetaldehyde consumption were determined by fitting time–concentration values to a first-order decay model. Butyaldehyde–DNPH was used to evaluate instrument performance.

Malondialdehyde

MDA in liver homogenate was assayed by TBARS Assay Kit (Cayman, MI) according to a modified manufacturer's protocol. In brief, the mixture of 25 μL liver homogenate, 25 μL SDS, and 1 mL color reagent was boiled at 99°C for 1 h. After cooling down on ice for 10 min, the sample was centrifuged at 1,600 g for 10 min and the absorbance was read at 540 nm by a spectrophotometer. MDA concentration was normalized by the amount of total protein determined by Pierce™ BCA Protein Assay Kit (Thermo Scientific).

Assessment of body weight, hemoglobin, locomotion, and skin pigmentation

To assess the effects of chronic ethanol challenge on total body mass, mice were weighed at pre-ethanol, 3, 6, 9, and 12 weeks time points.

Hemoglobin levels were determined in blood collected in EDTA tubes from the tail vein at pre-ethanol, 3, 6, 9, and 12 weeks time points and were automatically counted using the ADVIA 120 Hematology System (Siemens Healthineers, Erlangen, Germany).

The rotarod behavior test was used to evaluate mouse strength and coordination every 3 weeks during ethanol challenge. An automated four-lane rotarod unit (AccuScan Instruments, Columbus, OH) was used to evaluate locomotor activity. The apparatus has 7 cm diameter drums with grooves to improve grip. Drums were rotated at a fixed speed of 2 rotations/min (rpm) for the first 20 s, accelerated up to 30 rpm in the next 100 s, and then up to 60 rpm in the last 60 s. The time and rpm when mouse fell from or failed to walk on a drum were recorded. Failure to walk was defined as a mouse that did not fall from the drum, but clung in one position and went around twice. Tests were performed twice at each time point and the average rpm was calculated.

To assess skin pigmentation from chronic ethanol ingestion, pictures were taken of exposed ear, genitals, tail, and sole skin of each mouse. For analysis, darkness of the paw sole was quantified with ImageJ. All sole areas were traced and intensity of every dot in the traced area was measured as mean red–green–blue (RGB) intensity. Positive pigmentation was quantified as an intensity with mean RGB intensity of 90 or less after normalization to the mean RGB intensity of white background.

DNA damage and adducts

DNA damage was assessed by immunohistochemistry for γH2AX. γH2AX was stained with an antibody to phosphohistone H2A.X (Ser 139) (clone: 20E3, 1:100, Cell Signaling Technology, MA), and the number of γH2AX-positive cells and total epithelial cells on the basal membrane of the esophagus were counted using a 40 × objective. Measurement of the DNA adduct N2-et-dG in esophagus DNA was performed at the Proteomics Resource Center, Rockefeller University for n = 5 mice per group. DNA was extracted from a whole mouse esophagus using the QIAGEN (Germantown, MD) DNAeasy Blood & Tissue Kit, according to the manufacturer's protocol except with the addition of sodium cyanoborohydride (NaBH3CN) to each reagent of the kit (150 mM to cell lysis buffer and 100 mM to other reagents). Extracted DNA was dissolved in 60 μL of 10 mM Tris–HCl/5 mM EDTA buffer (pH 7.5), and the amount of DNA was measured by Nanodrop. Extracted DNA was mixed with 1.2 μL of 1 M citrate buffer (pH 6.0), 0.8 μL of 750 mM CaCl2, 45 U micrococcal nuclease (Worthington Biochemical Corporation, Lakewood, NJ), 0.15 U spleen phosphodiesterase (Worthington Biochemical Corporation), 4.8 μL of 8 M NaBH3CN, and 1 μL of 5 pm/μL internal standard. The mixture was incubated at 37°C for 3 h, and then 2.5 μL of 2 M Tris-HCl (pH 8.5), 2 μL of 45 mM ZnSO4, 6 U alkaline phosphatase (Sigma-Aldrich), 1 μL of 8 M NaBH3CN, and 1.5 μL distilled water were added. After incubation at 37°C for 3 h, nucleosides were extracted twice with 600 μL of chilled methanol and the combined supernatant was evaporated to dryness. The dried samples were resuspended in 100 μL LC/MS grade water (Optima™; Fisher Scientific), sonicated for 10 s, centrifuged at 4°C for 10 min, and supernatant was subjected to liquid chromatography tandem mass spectrometry (LC-MS/MS) analyses.

LC-MS/MS analyses were performed using a Vantage TSQ triple-stage quadrupole mass spectrometer (Thermo Fisher Scientific) equipped with a heated electrospray ionization source. The mass spectrometer was operated with the following parameters: positive ion polarity; spray voltage, 3,500 V; ion transfer capillary temperature, 300°C; source temperature, 250°C; sheath gas, 30 (arbitrary units); auxiliary gas, 10 (arbitrary units); collision gas, argon; and dwell time, 200 ms. The S-Lens voltage (S-lens) and collision energy (CE) were manually optimized for each MRM transition. The MRM transitions monitored were deoxyguanosine (dG), m/z 268 → 152 (CE = 12, S-lens = 50); [15N5]-N2-et-dG, m/z 301 → 185 (CE = 14, S-lens = 57); and N2-et-dG, m/z 296 → 180 (CE = 12, S-lens = 60). Chromatographic separation was performed on a Dionex Ultimate 3000 HPLC equipped with a Thermo Hypersil Gold aQ column (2.1 mm × 150 mm × 3 μm particle size). Column temperature was maintained at 36°C and the autosampler was set to 4°C. Mobile phase A consisted of 0.1% LC-grade formic acid (Pierce; Thermo Scientific) in water and mobile phase B consisted of 0.1% LC-grade formic acid in ACN. Separation was achieved using the following gradient (flow rate set at 0.4 mL/min): 0% B (0–6 min), 1% B (6–7.65 min), 6% B (7.65–9.35 min), held at 6% B (9.35–10 min), increased to 50% B (10–12 min), 75% B (12–14 min), held at 75% B (14–17 min), returning to 0% B (17–17.5 min), and re-equilibrating (17.5–30 min). The injection volume was 1–3 μL with ≥2 technical replicates per biological replicate. The amount of N2-et-dG in each DNA sample was determined from the ratio of the peak area of N2-et-dG relative to the internal standard [15N5] N2-et-dG and by using a calibration curve (0.6–2,500 fmol/μL, r2 ≥ 0.98). Owing to the high concentrations of dG in each DNA sample, the original resuspension stock was further diluted 100-fold followed by injection of 0.5–1 μL of this mixture onto the column. The amount of dG was then determined using a calibration curve (20–1,250 fmol/μL, r2 ≥ 0.97).

μCT and histological analysis of femurs

Femurs fixed in 4% paraformaldehyde then stored in 70% ethanol were scanned using a high-resolution Scanco μCT 35 (Scanco Medical AG, Bruttisellen, Switzerland). Specimens were scanned with an isotropic voxel size of 7 μm. For analysis of femoral bone mass, a region of trabecular bone 2.1 mm wide was contoured, starting 280 μm from the proximal end of the distal femoral growth plate. Femoral trabecular bone was thresholded at 211 permille and femoral cortical bone was thresholded at 350 permille. A Gaussian noise filter optimized for murine bone was applied to reduce noise in the thresholded two-dimensional (2D) image. Three-dimensional reconstructions were created by stacking the thresholded 2D images from the contoured regions.

For histological analysis, femurs were decalcified with 0.5 M EDTA (pH 8.0) and embedded in paraffin. Sections were deparaffinized and stained with hematoxylin and eosin. Metaphysis was defined as the area within 2 mm from the bottom of growth plate. Areas of metaphysis for each trabecular bone were traced and quantified by ImageJ. The ratio of trabecular bone in metaphysis was calculated by dividing the sum area of each trabecular bone in metaphysis by total area of metaphysis.

Statistical analysis

All data are presented as means ± standard error of the mean (SEM) unless otherwise stated; the “n” value for each group is stated in the figure or figure legend. Differences between groups were analyzed using an unpaired two-tailed Student's t-test. The behavior score and body temperature after ethanol exposure were also correlated with acetaldehyde levels. These correlations were evaluated using the correlation test for paired samples. r2 values >0.8 indicate a strong relationship between the test groups. p-Values <0.05 were considered significant for all comparisons.

Results

AAVrh.10hALDH2-mediated expression of hALDH2

Aldh2−/− and Aldh2E487K+/+ mice were treated with AAVrh.10hALDH2 (Supplementary Fig. S2) or AAVrh.10control (1011 gc) by intravenous administration. Sixteen weeks later, hALDH2 mRNA, protein levels, and ALDH2 enzymatic activity in liver were analyzed. AAVrh.10hALDH2-treated Aldh2−/− and Aldh2E487K+/+ mice had significantly higher hALDH2 mRNA expression in liver than AAVrh.10control-treated mice (Aldh2−/−, p < 10−4; Aldh2E487K+/+, p < 10−5; Fig. 1A). Western analysis revealed liver hALDH2 protein expression in AAVrh.10hALDH2-treated Aldh2−/− and Aldh2E487K+/+ mice (Aldh2−/−, p < 10−3; Aldh2E487K+/+, p < 10−2; Fig. 1B). Liver immunohistochemical staining of AAVrh.10hALDH2-treated Aldh2−/− and Aldh2E487K+/+ mice was consistent with the Western analysis. Liver hALDH2 positive cells were found mainly around hepatic and portal veins in AAVrh.10hALDH2-treated mice, whereas no positive cells were observed in AAVrh.10control-treated mice (Fig. 1C). In addition, AAVrh.10control-treated mice had undetectable levels of ALDH2 liver enzyme activity, whereas AAVrh.10hALDH2-treated mice showed high levels of enzymatic activity (Aldh2−/−, p < 10−4; Aldh2E487K+/+, p < 10−5) comparable with the wild-type C57Bl/6 mice (Aldh2−/−, p > 0.3; Aldh2E487K+/+, p > 0.4; Fig. 1D).

Figure 1. In vivo liver expression of human ALDH2 12 weeks after a single intravenous administration of AAVrh.10hALDH2 or AAVrh.10control (1011 gc) to Aldh2−/− and Aldh2E487K+/+ mice. C57Bl/6 mice were intravenously administered PBS. (A) Liver mRNA expression assessed by TaqMan RT-qPCR. (B) Liver protein expression assessed by Western analysis using an anti-HA tag antibody. GAPDH was used as a loading control. Quantification was assessed on the protein bands from the Western analysis. Two examples of Western analysis from each group are shown; each lane represents a different animal. (C) Liver hALDH2 immunohistochemistry assessed using an anti-HA antibody. (D) Liver ALDH2 enzymatic activity. Data are presented as means ± SEM. ALDH2, aldehyde dehydrogenase type 2; HA, hemagglutinin; PBS, phosphate-buffered saline; SEM, standard error of the mean. Color images are available online.

Water versus ethanol intake

Four weeks after vector administration, mice were supplied water or ethanol in their daily drinking bottles for 12 weeks (10% ethanol in water for the first 6 weeks and 15% ethanol in water for the second 6 weeks) ad libitum. Water or ethanol intake was measured every week. In all genotypes, water or ethanol intake increased proportionally to the number of mice in the cage (all r2 > 0.96, p < 10−9, Supplementary Fig. S3A). Ethanol intake was found to be significantly less than water intake for wild-type C57Bl/6, Aldh2−/− and Aldh2E487K+/+ mice (C57Bl/6, p < 10−4; Aldh2−/−, p < 10−8; Aldh2E487K+/+, p < 10−7; Supplementary Fig. S3A–C). In contrast, AAVrh.10hALDH2-treated Aldh2−/− and Aldh2E487K+/+ mice drank significantly more ethanol than AAVrh.10control-treated mice (Aldh2−/−, p < 10−5; Aldh2E487K+/+, p < 10−4; Supplementary Fig. S3B, C).

Systemic acetaldehyde and MDA levels after chronic ethanol exposure

To assess whether augmentation of ALDH2 enzyme levels in the liver by gene therapy could prevent the effects of chronic ethanol ingestion, Aldh2−/− and Aldh2E487K+/+ mice were administered AAVrh.10hALDH2 or AAVrh.10control intravenously 4 weeks before the addition of ethanol in the daily water supply. Serum acetaldehyde levels of AAVrh.10control-treated Aldh2−/− and Aldh2E487K+/+ mice given ethanol were significantly higher than those of mice given water for 12 weeks (Aldh2−/−, p < 10−2; Aldh2E487K+/+, p < 0.02; Fig. 2A). However, serum acetaldehyde levels in AAVrh.10hALDH2 vector-treated Aldh2−/− and Aldh2E487K+/+ mice given ethanol for 12 weeks were reduced to levels similar to the Aldh2−/− or Aldh2E487K+/+ mice given water and the wild-type C57Bl/6 mice given ethanol (all p > 0.7, Fig. 2A). In addition to metabolizing acetaldehyde, ALDH2 also processes reactive aldehydes derived from oxidative stress such as MDA, which is implicated in DNA adduct formation and mutagenesis.35,36 The level of MDA in the livers of AAVrh.10control-treated Aldh2−/− and Aldh2E487K+/+ mice given ethanol for 12 weeks was significantly higher than that of mice given water (Aldh2−/−, p < 10−3; Aldh2E487K+/+, p < 10−2; Fig. 2B). In contrast, the MDA level in AAVrh.10hALDH2-treated Aldh2−/− and Aldh2E487K+/+ mice was not significantly different from that of the ALDH2-deficient mice given water or the wild-type C57Bl/6 mice (all p > 0.1, Fig. 2B).

Figure 2. Effect of AAVrh.10hALDH2 therapy on serum acetaldehyde and liver MDA levels with chronic ethanol exposure. Aldh2−/− and Aldh2E487K+/+ mice were intravenously administered AAVrh.10hALDH2 (1011 gc) or AAVrh.10control (1011 gc). C57Bl/6 mice were intravenously administered PBS. Four weeks after vector administration, mice were challenged with water or ethanol for 12 weeks. (A) Serum acetaldehyde after 12 weeks chronic ethanol challenge. (B) MDA levels in liver after 12 weeks chronic ethanol exposure, normalized by the amount of total protein. Values are presented as means ± SEM. MDA, malondialdehyde. Color images are available online.

Body weight, hemoglobin, locomotion, and dermatological abnormalities

Prior studies have demonstrated that with chronic ethanol ingestion, Aldh2−/− and Aldh2E487K+/+ mice have decreased body weight and blood cell counts.9,37 To assess whether AAVrh.10hALDH2 therapy could prevent these effects of chronic ethanol consumption, mice were evaluated every 3 weeks for body weight, hemoglobin levels, and rotarod locomotion. Body weight and blood hemoglobin levels of AAVrh.10control-treated Aldh2−/− and Aldh2E487K+/+ mice decreased significantly for 12 weeks (Fig. 3A and Supplementary Fig. S4A, body weight, p < 10−3; Fig. 3B and Supplementary Fig. S4B, hemoglobin, p < 10−5). In parallel, these mice performed poorly on the rotarod test of locomotion (p < 10−2, Fig. 3C and Supplementary Fig. S4C). However, the body weight, hemoglobin levels, and rotarod performance for ALDH2-deficient mice treated with AAVrh.10hALDH2 were similar to those of the wild-type C57Bl/6 mice given ethanol for 12 weeks (Fig. 3 and Supplementary Fig. S4).

Figure 3. Effect of AAVrh.10hALDH2 therapy on body weight, hemoglobin levels, and locomotion (rotarod) during chronic ethanol exposure. Aldh2−/− and Aldh2E487K+/+ mice were intravenously administered AAVrh.10hALDH2 (1011 gc) or AAVrh.10control (1011 gc). C57Bl/6 mice were intravenously administered PBS. Four weeks after vector administration, mice were challenged with water or ethanol for 12 weeks. Tests were performed pre-exposure and 3, 6, 9, and 12 weeks during ethanol exposure. (A) Body weight; (B) hemoglobin; and (C) locomotion (rotarod) assessment of maximum rotations per minute without falling. Values are presented as means ± SEM. Color images are available online.

ALDH2*2 individuals as well as ALDH2-deficient mice develop skin hyperpigmentation with chronic ethanol

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