DETERMINATION OF THE FLAVIN-CONTAINING MONOOXYGENASE
mRNA LEVELS IN MOUSE LUNG AND LIVER
CHAPTER 1
INTRODUCTION
FMO Background
For many years, cytochrome P450s (CYPs) were thought to be the only enzymes that
catalyzed N- or S- oxygenation of nucleophilic heteroatom-containing small
molecules (Cashman, 1997). Not until another hepatic microsomal enzyme was
purified to homogeneity and demonstrated N-oxygenation of dimethylaniline in the
presence of NADPH were flavin-containing monooxygenases (FMOs) found to be
distinct from CYPs (Ziegler and Mitchell, 1972). Mammalian FMOs are a
xenobiotic-metabolizing gene family with each family having a single member that
binds FAD as a prosthetic group and NADPH as a cofactor; these groups and
molecular oxygen are required for biological activity. Like CYPs, FMOs are
localized to the endoplasmic reticulum and exhibit broad substrate specificity.
Substrates for FMO are structurally diverse compounds that contain a soft
nucleophile and commonly oxygenate xenobiotics containing nitrogen and sulfur
heteroatoms (Lee et al., 1980; Ziegler, 1984; Lomri et al., 1993a & b; Henderson
et al., 2004 a&b; recently reviewed by Krueger and Williams, 2005).
It is thought that the FMO gene family may have emerged from duplication of an
ancestral gene or through a series of independent gene duplication events (Hernandez
et al., 2004). The FMO genes are located within a cluster on chromosome one in
humans (Shephard et al., 1993; McCombie et al., 1996). Mammalian FMOs encode
1
protein products that are 532-558 amino acids in length and all contain highly
conserved FAD- and NADPH-binding sites (Atta-Asafo-Adjei et al., 1993; Lawton
and Philpot, 1993). In general, members of the mammalian Fmo gene family share
from 52-58% sequence identity (Hines et al., 1993; Lawton et al., 1994). Each, Fmo
family indicated by the first member (1, 2, etc.) has only one member. The sequence
identities of orthologous FMOs from different species are generally greater than 82%
(Lawton et al., 1990). Normally, orthologs retain the same function in the course of
evolution rendering the identification of orthologs critical for reliable prediction of
gene function in newly sequenced genomes. In humans, eleven FMO families exist
that include five pseudogenes; and in mouse nine families have been documented that
may all be expressed.
Human Ethnic Polymorphisms
Among the FMO families, considerable allelic variability in FMO enzyme function
can result from small base changes in the wild-type gene (Cashman et al., 2000;
Cashman 2004, Krueger and Williams, 2005). In particular, two important
polymorphisms exist in human FMO2. All Caucasians and Asians genotyped to date
are homologous for a genetic polymorphism, the FMO2*2 allele, in which a CT
transition at bp 1414 replaces a glutamine at amino acid residue 472 with a stop
codon (Dolphin et al., 1998; Whetsine et al., 2000). This change produces a nonfunctional truncated FMO2.2 protein from the FMO2*2 allele (Dolphin et al., 1998;
Krueger et al., 2002). FMO2.2 does not incorporate FAD and is not detected on
western blots of human pulmonary microsomes presumably due to degradation and
2
incorrect folding. The FMO2*1 allele, that encodes full-length functional FMO2.1
enzyme, is present in individuals from African (27%) and Hispanic
(2 to 7%) populations (Dolphin et al., 1998; Whetsine et al., 2000; Krueger et al.,
2002; Furnes et al., 2003; Krueger et al., 2004).
Potential for Differential Metabolism
Eleven distinct FMO genes exist in humans encoding five functional enzymes
(FMO1-5) and six pseudogenes. (Haining et al., 1997; Myers et al., 1997; Overby
et al., 1997; Whetstine et al., 2000; Yeung et al., 2000). Of the FMO isoforms,
FMO1, FMO2 and FMO3 are the main drug metabolizing enzymes in mammals and
all exhibit broad substrate specificities. For example, FMO1 appears to have a less
restricted substrate access channel and accepts larger N- and S-containing
nucleophiles than human FMO3 (Taylor and Ziegler, 1987; Hvattum et al., 1991;
Attar et al., 2003; Lomri et al., 1993b; Overby et al., 1995); however, both show
efficient activity for S-oxygenation of sulfur-containing chemicals, such as thioureas
and thioamides, similar to FMO2.
In general, metabolism by FMO yields metabolic products that are more polar and
less toxic or less biologically active and more readily excreted than the parent
compound (Ziegler, 1984; Damani, 1988). This function allows the body to remove
unwanted natural products from the diet (e.g., amine-, sulfide-, phosphorus-, and
other nucleophilic heteroatom-containing compounds). However, in contrast to other
heteroatoms, sulfur oxygenation sometimes results in bioactivation (Ziegler, 1991;
3
Cashman, 1995; Genter et al., 1995). FMO2 S-oxygenation is important because of
the potential toxicities to lung. For example, α-naphthylthiourea (ANTU) is a
rodenticide that is a sulfur-containing substrate and when ANTU is S-oxygenated it
can covalently bind to proteins and deplete glutathione (GSH), an antioxidant
involved in detoxification and elimination of toxic materials. This depletion produces
toxicity that can lead to severe pulmonary edema and vascular injury. (Scott et al.,
1990; Boyd and Neal, 1976; Lee et al., 1980; Hardwick et al., 1991; Barton, et al.,
2000). A major metabolite observed by HPLC following incubation of ANTU with
Sf9 expressed FMO2.1 is the sulfenic acid (Henderson et al., 2004b). This is
important because ANTU is a good substrate for FMOs, including lung FMO2. The
sulfenic acid can be further oxidized to sulfinic acid or combined with GSH
(Henderson et al., 2004b) with the net result being conversion of GSH to oxidized
GSH (GSSG) and regeneration of ANTU that establishes a redox cycle which can
lead to oxidative stress and toxicity.
The demonstration that FMO2.1 functions in the in vitro bioactivation of thioureas
such as ethylenethiourea and ANTU (Henderson et al., 2004b) and in the potential
detoxification of organophosphate insecticides such as phorate and disulfoton
(Henderson et al., 2004a) is significant. These findings could have important
implications for Hispanic and African-American individuals with the functional
(FMO2*1) allele. They may exhibit distinct outcomes compared with the general
population following environmental or occupational exposures to such chemicals or in
therapeutic response to drugs that are substrates for FMO2.1.
4
Utility of the Knock-out Mouse Model
Generally, knock-out mouse models are used to elucidate gene functions, model
genetic disorders, and assist in drug development. In a knockout mouse, gene
function is destroyed by elimination of an essential part of the gene of interest. The
Fmo2 knock-out mouse model under development in this laboratory will be used to
reveal the role of human pulmonary FMO2 in xenobiotic metabolism and toxicity. In
the Fmo2 knockout mouse the null Fmo2 allele will represent the human population
that contains the non-functional allele (FMO2*2). The wild-type mouse containing
the functional Fmo2 allele will be a model for the minority population that has the
FMO2*1 allele encoding functional protein. Using the knock-out mouse models, the
role of active FMO2 in metabolism of foreign substrates can be discovered.
Dietary Modulation of FMO
The comparison of flavin-containing monooxygenase levels in liver of adult male
Sprague-Dawley rats fed total parental (TPN) and “commercial rat chow” (Purinabrand rat chow) for seven days showed a reduction of FMO protein that included
N,N-dimethylaniline N-oxidase activity to 34% of the activity in chow-fed controls.
TPN diet contains semisynthetic ingredients including 27% dextrose, 5% amino acids
and supplemental vitamins and minerals (Kaderlik et al., 1991). Interestingly, similar
results were demonstrated in rats orally fed with chemically defined, semisynthetic
diets similar to TPN. Rats fed these semisythetic diets for 3-4 days demonstrated
lower liver FMO activity. This loss of function; however, was reversible since FMO
activity could be increased to near control levels when chow feeding was resumed or
5
when the semisynthetic diets were supplemented with chow extract (Kaderlik et al.,
1991).
There are patients diagnosed with trimethylaminuria who benefit from knowing the
factors that affect FMO activity. In Cashman et al., (2004), rats fed a TPN + choline
diet displayed a 3-fold increase in hepatic microsomal FMO with FMO1 protein
decreased 1.4-fold and FMO3 increased 1.3-fold. This is important because choline
is a precursor for trimethylamine (TMA) and in the absence of FMO3, TMA cannot
be N-oxygenated to its detoxification product. This has been shown to cause
trimethylaminuria or “fish odor syndrome” where individuals lacking FMO3 smell
like fish. Understanding that a diet of TPN + choline can increase the FMO3 activity
levels has beneficial medical implications for these patients.
There is further experimental evidence that diet regulates FMO expression. Nnane
and Damani, (2001) studied the pharmacokinetic profile of the FMO substrate
trimethylamine (TMA) in male Wistar rats fed either a synthetic diet or chow-based
diet. Male Wistar rats fed a synthetic diet had a two-fold decrease in the clearance of
TMA compared with the chow-based diet. Concentrations of the N-oxide metabolite
in blood after intravenous administration of TMA peaked at 1.25 h in rats placed on
the synthetic diet, as opposed to 0.75 h in rats placed on normal laboratory rat chow.
FMO3 is involved in the oxygenation of TMA and so the altered pharmacokinetic
profile of TMA and its N-oxide suggest a reduced drug-elimination capacity in rats
placed on the synthetic diet presumably due to reduced FMO3 levels. Another study
6
that supports dietary modulation of FMO is the demonstration that dietary indole-3carbinol (a naturally occurring cruciferous vegetable or plant alkaloid) reduces FMO
expression and activity in rat liver and intestine (Larsen-Su and Williams, 1996).
Sex, Development, and Tissue Specific Expression of FMO2
The FMO gene family exhibits sex, development, and tissue specific expression.
This has been characterized in a number of animal species including humans, mice,
rats, and rabbits (Hines et al., 1994) using methods such as RNase protection assays
and qPCR to quantify mRNA (Janmohamed et al., 2001; 2004); and western blot
analysis (Koukouritaki et al., 2002) and microsomal activity assays (Overby et al.,
1995; Shehin-Johnson et al., 1994) to detect protein level and activity. Most studies
have focused on liver and kidney tissues with high capacity for phase I drug
metabolism.
Sex-specific expression of Fmo in mouse liver was documented by Falls et al.,
(1995). Their studies showed that: Fmo mRNA levels in female were two to three
times greater than that in male and Fmo1 expression is equivalent to the expression of
Fmo3 in female with Fmo2 not detected. In males, Fmo2 and Fmo3 mRNA were
undetectable. Fmo3 was shown to increase with age, Fmo3 protein levels were twice
that of Fmo1 and mRNA levels were four times greater than Fmo1. Fmo1 was the
primary Fmo detected in fetus whereas Fmo3 was the main Fmo found in adult mice.
In contrast, patterns of mouse Fmo expression documented by Janmohamed et al.,
(2004), using RNase protection assays to quantitate total RNA, showed that Fmo
mRNA levels in liver of 129/SV and C57BL6J female mice were Fmo3 twice that of
7
Fmo1; Fmo1 levels were generally detected in males and Fmo3 was detected at 3 wk
of age.
Fmo2 expression in mouse lung remains somewhat uncertain. Studies performed by
Falls et al., (1995) did not include lung; however, mouse studies performed by
Janmohamed et al., (2004), demonstrated high levels of Fmo1, and low levels of
Fmo2 and Fmo3 in lung. This is in contrast to our unpublished preliminary work that
shows Fmo2 is abundant in lung, Fmo3 mRNA levels are moderate and Fmo1 levels
are low (see Figure 1 and Table 1). Studies show that FMO2 is the sole FMO in the
lung of most mammals (Lawton et al., 1990; Nikbakht et al., 1992; Yueh et al.,
1997). Mouse strain (129/SV vs. 129/SV and C57BL6J F1hybrid ), age (8 weeks vs.
12 weeks), diet (Harlan Teklad TRM vs. AIN93G), and method differences (RNase
protection assay vs. qPCR, primers for reverse transcription and detection and
normalization gene) are also possible causes of this inter-laboratory discrepancy.
Development-specific expression of FMO2 has been demonstrated in fetal and
neonatal rabbit lung (Larsen-Su et al., 1998). During late gestation, FMO2 levels in
rabbit fetal lung were high. However, at parturition, FMO2 levels fell dramatically
and recovered 21 days post-parturition. The development-specific expression of
FMO2 suggests that FMOs may have the potential to play a significant role in the
metabolism of endogenous compounds as well as the detoxification of foreign
substances.
8
In addition to sex-specific expression of FMO suggesting sex-steroid hormonal
regulation, it is demonstrated in Rouer et al., (1987) that microsomal FMO activation
in the liver of mice occurs in the pathological states that include type 1 and type 2
diabetes. The formation of impiramine N-oxide is increased with diabetes in mice.
This suggested that flavin-containing monooxgenase could be controlled by
glycoregulatory hormones besided sex-steroid hormonal regulation as implied in the
sex-dependent expression of FMO.
Tissue specific expression of FMO exists. Human FMO1 mRNA is the most
prevalent FMO in adult kidney (reviewed by Cashman and Zhang, 2006) and FMO2
is the dominant isoform in the adult mammalian and human lung (Lawton et al.,
1990; Nikbakht et al., 1992; Yueh et al., 1997). Unique to humans, FMO3 is the
most abundant FMO family member in the adult liver, whereas FMO1 dominates in
most animal models except for female mouse models where Fmo3 is more abundant.
Early studies documented that FMO1 is the most abundant FMO enzyme in the
human fetal liver, whereas FMO3 is essentially absent in the fetal liver (Hines and
McCarver, 2001). The timing of the FMO1/FMO3 switch or potential control
mechanisms are documented in Koukouritaki et al., (2001). Using microsomal
fractions from 240 human liver samples and a western blot method, FMO1 protein
expression was found to be more abundant in the embryo (8-5 wk gestation; 7.8 +/5.3 pmol/mg) with low levels of FMO3. FMO1 was generally suppressed within 3 d
postpartum, but not during gestational age, and FMO3 was not expressed during the
neonatal period; however, was detected in individuals by 1-2 y of age at intermediate
9
levels until 11 y (12.7 +/- 8.0 pmol/mg protein). Furthermore, in individuals between
the ages of 11 to 18 y of age, a gender-independent increase in FMO3 expression was
observed (Hines et al., 2002). Other studies showed that the FMO5 mRNA level was
nearly as abundant as FMO3 in human adult liver (reviewed by Cashman and Zhang,
2006).
Statement of Purpose
The determination of Fmo2 in mouse lung is of interest because of its potential role in
the development of a knockout mouse model that should elucidate the role of FMO2.1
in the metabolism of environmental chemicals for which lung is a target or route of
administration. If Fmo1 and Fmo3 are significantly expressed then the utility of the
knock-out mouse model will be reduced because of overlap in substrate specificity.
High levels of Fmo1, and low levels of Fmo2 and Fmo3 in mouse lung were recently
reported (Janmohamed et al., 2004), in contrast to our own preliminary findings that
Fmo2 was abundant in lung, Fmo3 message levels were moderate and Fmo1 levels
were low (Figure 1, Table 1)
10
0.06
0.05
pg Fmo
pg Fmo/pg actin
FMO2
FMO1
0.04
FMO3
0.03
0.02
0.01
0
female lung
male lung
Figure 1. Mean expression of Fmo isoforms (pg Fmo/pg actin) in lung of mice fed
AIN93G. Each bar shows the mean and standard deviation (s.d.) of three mice.
Table 1. Mean expression of Fmo isoforms (pg Fmo/pg actin) and s.d. in lung of mice
fed AIN93G for nine weeks.
Sex
Female
Male
FMO2
mean
s.d.
0.0468 0.0070
0.0485 0.0054
FMO1
mean
0.0053
0.0081
s.d.
0.0003
0.0011
FMO3
Mean
s.d.
0.0097
0.0011
0.0096
0.0011
Possible causes of this inter-laboratory discrepancy include mouse strain (129/SV vs.
129/SV C57BL6J F1 hybrid), age (8 weeks vs. 12 weeks), and diet (Harlan Teklad
TRM vs. AIN93G). This study was designed to determine whether dietary differences
would explain observed differences in Fmo expression since dietary modulation has
been documented. Determining the levels of Fmo in mouse lung and liver will assist in
the best utilization of the knock-out mouse currently under development that should
elucidate the role of human pulmonary FMO2 in xenobiotic metabolism and toxicity.
The research questions are as follows:
11
(1) Will diet modulate Fmo mRNA expression?
(2) Will Fmo2 be the predominant Fmo isoform expressed in the lung of mice fed
AIN93G diet?
Our working hypothesis was that the AIN93G (semi-synthetic) and NIH31 (chowbased) diets modulate expression of Fmo in lung, and that Fmo2 would be the
predominant isoform expressed in lung of mice fed AIN93G. A correct hypothesis
would indicate the most appropriate diet that would be critical during studies with
Fmo2 knock-out mice. An Fmo2 knock-out mouse will represent the majority of the
human population that expresses the non-functional (FMO2.2) protein, whereas, the
wild-type mice that produce functional Fmo2 protein will represent those humans
producing functional FMO2.1.
Method of Investigation
In this study, we used a highly sensitive qPCR method to determine the levels and
distribution of Fmo using mRNA. The nature of the quantitative analysis allowed a
comparison among Fmo1, Fmo2, Fmo3 normalized by the levels of actin
(a housekeeping gene) detected in male and female mice on the chow-based or
synthetic diet. Other methodologies have been suggested such as enzyme assays;
however, this is not feasible due to broad, overlapping substrate specificities among
Fmo1, Fmo2 and Fmo3. Western detection was also not the method of choice because
the polyclonal antibodies that we currently use cross react with each of the other
isoforms and the isoforms could not be resolved by SDS-PAGE. This antibody crossreaction leads to inaccurate measurement or estimation of Fmo isoform levels.
12
Principal Results and Conclusions
Our working hypothesis was that the AIN93G (semi-synthetic) and NIH31 (chowbased) diets modulate expression of FMO in lung; and that in mice fed AIN93G, Fmo2
would be the predominant isoform expressed. Patterns observed were greater
abundance of Fmo1 followed by Fmo3 and low detection of Fmo2 in lung and liver,
regardless of diet. However, due to the high variability found in the data further studies
are required. Additional studies to examine the influence of strain and age differences
on Fmo expression are still desirable. This paper will discuss findings of whether
dietary modulation of AIN93G and NIH31 exist in Fmo expression and implications
from the patterns demonstrating greater Fmo1 levels as compared to Fmo2 regardless
of diet.
13
CHAPTER 2
METHODS AND MATERIALS
Chemicals
TRIzol and Super Script® III First-Strand Synthesis SuperMix for qPCR were
purchased from Invitrogen (Carlsbad, CA). Isopropyl alcohol was from LabGuard
(Philipsberg, NJ). Ethanol was from Aaper Alcohol (Shelbyville, KY). Distilled
RNase and DNase free H2O was from Gibco (Grand Island, NY). Tris-HCL pH 7.5
was from USB (Cleveland, OH). ß-mercaptoethanol was from Sigma (St. Louis, MO).
The RNeasy mini kit was from Qiagen (Valencia, CA). DyNamo SYBR Green qPCR
kit was from Finnzymes (Espoo, Finland).
RNA and Tissue Samples
Eight 129S1/SV/IMJ male (11.8-16.0g) and eight female (12.0-15.4g) mice were
obtained at three weeks of age from Jackson Laboratory (Bar Harbor, ME). This study
was approved by the Institutional Animal Care and Use Committee, IACUC (Corvallis,
OR). The animals were maintained in a controlled environment of constant room
temperature (22-25°C) and 12-h light cycle. Animal health was monitored throughout
the study by determining changes in body weight (see Appendix Figure A1), behavior
and facial/skin color and appearance. The eight male and eight female mice were each
divided into two groups. Half of each sex were maintained on a general laboratory
chow-based diet, NIH31, (HarlanTeklad, Madison, WI) while the balance were
assigned a semi-synthetic diet, AIN93G, (MP Biomedicals, Irvine, CA) ad libitum at
the Laboratory for Animal Research Center (LARC) for five weeks. The composition
of the diets is detailed in Table A1 of the appendix. The mice were euthanized at eight
14
weeks of age by exposure to carbon dioxide. The tissues were harvested and collected
using protocols approved by IACUC. Total RNA from lung and liver of mice was
prepared from 50-100 mg of tissue with TRIzol reagent using a protocol from the
manufacturer. RNA purity and quantity of samples were checked using an Agilent
2100 Bionanalyzer (Palo Alto, CA) and the NanoDrop Technologies NanoDrop
Spectrophotometer (Wilmington, DE). RNA samples were stored at -80°C until
analysis.
Reverse Transcription
Lung and liver RNA (2 µg) were treated with deoxyribonuclease I, amplification grade
(Invitrogen) following manufacturers protocol. This was to eliminate any genomic
DNA contamination before making cDNA. First strand synthesis of cDNA was done
with Super Script® III First-Strand Synthesis SuperMix for qPCR (Invitrogen). A total
reaction volume of 40 µl (0.1 µM muF1-63r, muF2-74r and muF3-69r, 1.85 µM oligo
dT18, 0.5 mM dNTP and H2O, 1X DNase buffer, 1.8 µl DNase, 5.0 µM dithiothreitol,
80 µl RNase Out, 400 U Superscript III) was used.
Table 2. q-PCR primers and conditions
Primer
Name
act4-2
act-6r
muF1-63r
muF1-149
muF1-150r
muF2-73
muF2-74r
muF3-69r
Gene
actin
actin
Fmo1
Fmo1
Fmo1
Fmo2
Fmo2
Fmo3
Sequence from 5' to 3'
TCC CTG GAG AAG AGC TAC GAG C
GCT TGC TGA TCC ACA TCT GCT G
TTA CAG GAA AAT CAA GAA GAC AGC T
TGT CTC TGG ACA GTG GGA AGT
CAT TCC AAC TAC AAG GAC TCG
AGG CTC CAT CTT CCC AAC CGT A
CCG GGT CTT TAA GGG TTT CAG G
TCA GAT CAA CAC AAG GAA CAA AGC
Usage
1
1
2
1
1
1
1,2
2
15
muF3-151
muF3-152r
Fmo3 CAC CAC TGA AAA GCA CGG TA
Fmo3 GTT TAA AGG CAC CAA ACC ATA G
1
1
Table 2. Continued
Primer
Name
act4-2
act6r
muF1-63r
muF1-149
muF1-150r
muF2-73
muF2-74r
muF3-69r
muF3-151
muF3-152r
Exon
bpLocation cDNA
4&6
376
4&6
376
9
4&5
217
4&5
217
7&8
382
7&8
382
9
4&6
445
4&6
445
All reactions were run in a MJ Research Thermocycler, in groups of ten, on separate
days, for 1 h at 50°C prior to being inactivated for 5 min at 70°C. The 40 µl reactions
were brought to 100 µl with nuclease-free water. The cDNAs were stored at 4°C until
further evaluation was performed.
qPCR Conditions
Gene-specific primers were designed for mouse Fmo1, Fmo2, Fmo3 and actin for
qPCR (Table 2). The primer pairs amplified products across at least one intron so that
any contamination by genomic DNA could be detected. All qPCR reactions were run
in 0.2 ml PCR tubes using the MJ Research Opticon Monitor 3 BioRad Laboratories
(Hercules, CA). Actin was selected as the housekeeping gene because it is assumed to
be expressed consistently in mammalian tissues and has been widely used for
normalization. Actin cDNA was a gift from Dr. Eric Andreasen (Oregon State
16
University). The clone consisted of a partial fragment of the zebrafish ß-actin gene that
was cloned into an expression vector and was used as the template for the standard
curve. The qPCR reactions were prepared as 20 µl reactions using the following
components: primer mix (0.1 µM, each reverse and forward gene specific primers), 1 µl
cDNA and 1X DyNAmo SYBR Green I. The qPCR conditions were optimized by
temperature gradients and melting curve analysis using varying annealing temperature
and primer pairs, and evaluated on NuPAGE TBE 6% acrylamide gel (Invitrogen)
electrophoresis to verify a single amplified product. An annealing temperature of 55°C
(Tm) was selected for all reactions. The qPCR reactions were held at 95°C for 10 min
followed by 30 cycles at 94°C for 15 s, 55°C for 30 s, and 72°C for 30 s. This was
followed by melting curve data collection and analysis. For each plate, standards and
samples were set-up in duplicates.
Data Normalization and Gene Quantification
Fmo standards were made from gel purified Fmo products amplified by PCR and then
quantitated to make a dilution curve. The Fmo products were purified with Gene Clean
(Qbiogene, Morgan Irvine, CA) using the provided protocol Picogreen assay
(Invitrogen, Carlsbad, CA) was used to quantitate Fmo standards. Standards for mouse
Fmo1, Fmo2 and Fmo3 and zebrafish actin with TE buffer (10 mM Tris, 1 mM EDTA
pH 7.5) were serially diluted to desired values from 10-5 to 102 pg/µl to generate
standard calibration curves for qPCR analysis. Using the Opticon Monitor 3 Software,
standard curves was made using dilutions of known quantities of Fmo standard
templates and actin by setting a cycle threshold level at the linear phase of the reaction
17
on the log scale of fluorescence of Syber Green Dye. This was done for all standards
and unknowns in a run and was used to quantitate unknown samples.
Melting curve analysis was made at reaction completion as an additional check to
ensure that a single product was being formed by the PCR reaction. Additional peaks
or shoulders indicated that there is some non-specific priming and possibly primer
dimer formation contributing to the fluorescence signal and distorting the quantitation.
The software calculated pg of Fmo1, Fmo2, Fmo3 and actin in the samples based on a
standard curve method. A normalization factor was then calculated based on the
quantities of Fmo1, Fmo2, Fmo3 and actin generated from the standard curve
(Fmo pg/actin pg) in order to compare the expression level of Fmo1, Fmo2 and Fmo3
in lung and liver. Fmo1, Fmo2 and Fmo3 quantities were normalized with actin
because actin assuming to be uniform actin expression in each tissue would normalize
for any variability in the amount of RNA used.
18
CHAPTER 3
RESULTS
Primer Test
Original primer pairs used for Fmo1 and Fmo3 did not produce a single clean peak (not
shown); thus, alternate primers were used for qPCR (see Figure 2a-d) that produce
cleaner products for mouse Fmo1 and mouse Fmo3 in melting curve analysis than the
original primer pairs did. NuPAGE TBE gels, 6% acrylamide, were used to determine
product homogeneity not shown. The original mouse Fmo2 reverse and forward
primers (muF2-73 and muF2-74r) and actin (act4-2 and act-6r) were suitable without
modification.
a.
b.
c.
d.
Figure 2. Primer test using Fmo and actin-specific primers with gene-specific cDNA
dilution. Melting curves for (a) Fmo2 with muF2-73 and muF2-74r primers. (b) actin
with act4-2, act-6r primers. (c) Fmo3 with muF3-151 and muF3-152r primers. (d)
Fmo1 with muF1-149 and muF1-150r primers.
19
Fmo Expression
Fmo expression levels were determined by qPCR in male and female mice fed either
AIN93G or NIH31. Figure 3 & 4, summarize findings for actin-normalized mRNA
levels of Fmo1, Fmo2 and Fmo3 expressed in lung tissue as a function of diet by sex.
The data generated by qPCR show Fmo1 to be the most highly expressed isoform in
mouse lung regardless of diet. Patterns show possible sex-specific expression of Fmo
levels with males expressing two to three times greater Fmo1 levels than females. As
for liver tissue, a sex-specific expression pattern of Fmo levels seems apparent. Fmo1
is the more highly expressed isoform in male mice, whereas Fmo3 is the most abundant
isoform expressed in female mice on both diets as shown in Figure 4. Actin levels liver
were not equal to actin levels in lung; therefore, comparisons of Fmo levels are only
possible within a tissue. Expression levels were not statistically analyzed due to the
high variability observed.
20
pg Fmo/ 2 ng RNA/pg actin
8.5
8.0
7.5
7.0
6.5
6.0
5.5
5.0
4.5
4.0
3.5
3.0
2.5
2.0
1.5
1.0
0.5
0.0
Fmo2
Fmo1
Fmo3
NIH-31
NIH-31
Male
Female
1
AIN93-G
AIN93-G
Male
Female
Figure 3. Mean expression of Fmo isoforms (pg/2ng mRNA/pg actin) in lung of mice
fed two diets. Each bar shows the mean and s.d. of four mice.
24.0
pg Fmo/ 2 ng RNA/pg actin
22.0
20.0
Fmo2
18.0
Fmo1
16.0
Fmo3
14.0
12.0
10.0
8.0
6.0
4.0
2.0
0.0
NIH31
NIH31
Male
Female
1
AIN93G
AIN93G
Male
Female
Figure 4. Mean expression of Fmo isoforms (pg/2ng mRNA/pg actin) in liver of mice
fed two diets. Each bar shows the mean and s.d. of four mice.
21
Dietary Modulation of Fmo
Fmo1, Fmo2 and Fmo3 were quantitated using qPCR to discover if the synthetic and
chow-based diets modulate Fmo isoform expression at the mRNA level. Figures 3 and
4 visualize the average Fmo isoform levels and standard deviations in each diet and are
separated by sex. Patterns observed show that diet may make a difference in the
expression level of some Fmos.
In lung tissue of mice, patterns showed that females fed the NIH31 (chow-based) diet
expressed twice the Fmo3 levels compared to those fed AIN93G (semi-pure, synthetic)
diet. Male mice fed the AIN93G diet appeared to express greater levels of Fmo2;
whereas female mice fed the NIH31 diet showed patterns of greater Fmo2 expression.
Furthermore, on average, male mice on the AIN93G diet expressed more Fmo1 than
those fed the NIH31 diet.
As for liver tissue of mice, dietary modulation of Fmo levels may also exist. Females
on the chow-based NIH31 diet exhibited lower levels of Fmo3 expression as compared
to those fed the semi-pure, synthetic AIN93G diet. In addition, male mice fed AIN93G
expressed Fmo3; whereas those on the NIH31 diet did not detect Fmo3.
Nonetheless,
it is clear that, in contrast to our earlier results, Fmo2 is not the most abundant message
in any tissue.
22
CHAPTER 4
DISCUSSION
Observations and Limitations
We had hypothesized that we would demonstrate differential dietary regulation by
AIN93G (semi-pure, synthetic) and NIH31 (chow-based) diets because these diets have
been show to modulate FMO levels in other studies. Nnane and Damani, (2001)
demonstrated that male rats fed a synthetic diet vs. chow based exhibited a 2-fold
decrease in the clearance of TMA. This suggested a reduced drug-elimination capacity
in rat placed on the synthetic diet presumably due to reduced FMO3 levels. Another
study that supports dietary modulation of FMO is the demonstration that dietary indole3-carbinol (a naturally occurring cruciferous vegetable or plant alkaloid) reduces Fmo
expression and activity in rat liver and intestine (Larsen-Su and Williams, 1996).
The results of this study do not reject the hypotheses that diet modulates Fmo
expression. However, it is clear in this study that on either diet Fmo2 is not the
predominant isoform for mice fed in this study in contrast to our preliminary findings.
Observed patterns show that Fmo1 mRNA is more abundant than Fmo2 on either diet.
Though these patterns were observed, high variations in the data obscure dietary
differences.
The patterns displayed in this study do not correlate well with other work that has been
published on Fmo isoform expression in mouse liver. In the study by Falls et al.,
(1995), Fmo levels in female mouse liver were found to be two to three times greater
23
than in male mouse liver and Fmo1 and Fmo3 message levels were reported to be
equivalent with lower expression of Fmo2 (Fmo1~Fmo3>>Fmo2). Male mice in their
study had twice the Fmo1 expression compared to Fmo3 and equivalent Fmo3 and
Fmo2 expression. These patterns do exist in this study with the exception that Fmo3 is
expressed in liver of male mice fed AIN93G. Similarly, patterns of Fmo expression in
the Janmohamed et al., study (2004) show that Fmo mRNA levels in liver of 129/SV
female mice fed a chow-based diet were Fmo3 twice that of Fmo1 and only Fmo1
levels were detected in males using RNase protection assays to quantitate total RNA
isolated. These findings correlate also correlate with the results in this study with the
exception of that Fmo3 is expressed in liver of male mice fed AIN93G
The lung expression results we documented in this study slightly correlated with the
Janmohamed et al., (2004) study. Their work found that Fmo mRNA levels in lung of
male and female mice were Fmo1 is three times more expressed than Fmo3, and Fmo3
and Fmo2 are equivalent in expression. In this study, lung of female mice showed
patterns demonstrating Fmo1 levels that were twice that Fmo3 and Fmo3 slightly more
expressed than Fmo2. The levels in male were Fmo1 three times more expressed than
Fmo3 with Fmo3 and Fmo2 equivalent in expression. The differences in the
Janmohamed et al., study and this work is that Fmo3 and Fmo2 levels are not
equivalent in females.
The high variation in the data could have resulted from several sources including
variation that exists naturally in animal tissues, RNA degradation, variations in RNA
24
quantity used to make first strand cDNA and primers used to synthesize first strand
cDNA, etc. Another factor could be that this study used isoform-specific primers
(0.1 μM muF1-63r and muF3-69r) for the transcription reactions that were not ideal.
These primers were part of initial primer pairs that did not yield clean PCR products, so
transcription may not have been uniform between isoforms. In addition, the stability of
cDNA synthesis components to freeze-thaw cycles and not making all cDNA at the
same time could have contributed to the high variation.
RNA integrity and purity are not believed to have contributed to the study variability.
We evaluated total RNA quality using the Agilent 2100 Bioanalyzer. All samples had
acceptable RNA Integrity Numbers, values in the range of 7.5-10. In addition,
electropherogram summaries indicated that 28s:18s rRNA ratios ranged between1.8 to
2.0. This indicated that RNA was not degraded (Appendix Figure A2).
Recommendations
The high variation in this data could be reduced by using a larger sample size. In this
study, the sample size was four. According to statistical models, using larger sample
size will lessen the average variation. In addition, a new housekeeping gene should be
used. The actin gene in this study had high variability among same tissue samples and
was not present and a constant level in liver and lung. Testing for the appropriate
housekeeping gene may alleviate the high variability in this study. Technique was
another issue and so pipetting errors, and processing first strand synthesis reactions at
different time intervals could have contributed to degradation of components and RNA
25
degradation since differences in DNase treatment time and EDTA inactivation can
occur. To avoid this problem, processing all liver samples or lung tissues at one time,
to ensure consistency, is recommended.
Furthermore, additional research on the possible influence of strain and age (extending
the time period) differences on Fmo expression will need be conducted to identify an
appropriate strain compatible with Fmo2 knock-out studies. Our preliminary
unpublished studies employed a hybrid mouse strain of 129/SV and C57BL6J, whereas
the Janmohamed et al., study (2004) used distinct 129/SV and C57BL6J strains. It is
possible that the hybrid strain causes Fmo expression differences that would not be
predicted from the parent strains from different sources. Understanding whether strain
and age modulate Fmo expression will help to identify with the parental strain for the
Fmo2 knock-out mouse model. The knock-out model, in turn, should provide
important information regarding the responses to various environmental pulmonary
toxicants by 27% of African Americans and 5% of Hispanics carrying the FMO2*1
allele.
Ongoing Studies
Mouse strain differences and their influence on Fmo levels are currently in progress. This
study is using female mice of eight strains fed chow-based diets until eight weeks of age.
Random primers were used for first strand synthesis and gene specific primers, identical to
the ones used in this study, for qPCR. Three housekeeping genes are being utilized: the
RPL13A (large ribosomal protein), HPRT (hypoxanthinephophoribosyltransferase) and
26
TBP (TATA-box binding protein). To date, preliminary results indicate that there is Fmo1
and Fmo2 expression lung tissue from each strain with Fmo2 greater than or equal to
Fmo1, and Fmo3 not detected. This was observed for 129 S/V strain normalized with
HPRT results and is different from the Fmo levels detected in our dietary study
(Fmo1>>Fmo3~Fmo2) reported here. Fmo1 and Fmo2 expression levels may vary by
strain. C57BL6J mice have lower average levels of Fmo1 and Fmo2, whereas 129/SV
appears to have a greater abundance of both isoforms.
27
CHAPTER 5
REFERENCES
Atta-Asafo-Adjei E., Lawton M.P., and Philpot R.M. (1993). Cloning, sequencing,
distribution, and expression in Escherichia coli of flavin-containing monooxygenase
1C1. Evidence for a third gene subfamily in rabbits. J Biol Chem, 268, 9681-9689.
Attar M., Dong D., Ling K.H. and Tang-Liu D.D. (2003). Cytochrome P450 2C8 and
flavin-containing monooxygenases are involved in the metabolism of tazarotenic acid
in humans. Drug Metab Dispos, 31, 476-481.
Barton C.C., Bucci T.J., Lomax L.G., Warbritton A.G., and Mehendale H.M. (2000).
Stimulated pulmonary cell hyperplasia underlies resistance to α-Naphylthiourea.
Toxicology, 143, 167-181.
Boyd M.R. and Neal R.A. (1976). Studies on the mechanism of toxicity and of
development of tolerance to the pulmonary toxin, alpha-naphthylthiourea (ANTU).
Drug Metab Dispos, 4, 314-322.
Cashman J.R. (1995) Structural and catalytic properties of the mammalian flavincontaining monooxygenase. Chem Res Toxicol, 8, 166-181.
Cashman J.R. (1997). Monoamine oxidase and flavin-containing monooxygenase.
Comprehensive Toxicology. (Guengerich F.P. ed), 3. Elsevier Science, New York, NY.
Cashman J.R., Akerman B.R., Forrest S.M. and Treacy E.P. (2000). Population-specific
polymorphisms of the human FMO3 gene: significance for detoxification. Drug Metab
Dispos, 28, 169-173.
Cashman J.R., Lattard V. and Lin J. (2004). Effect of total parental nutrition and
choline on hepatic flavin-containing and cytochrome P450 monooxgenase activity in
rats. Drug Metab Dispos, 32, 222-229.
Cashman J.R. (2004). The implications of polymorphism in mammalian flavincontaining monooxygenases in drug discovery and development. Drug Discovery
Today, 9.
Cashman J.R., and Zhang J. (2006). Human flavin-containing monooxygenases. Annu
Rev Pharmacol Toxicol, 46, 65-100.
Damani L.A. (1988). The flavin-containing monooxygenase as an amine oxidase, in
Metabolism of Xenobiotics. (Gorrod J.W., Oelschlaeger H. and Caldwell J. eds), 59-70,
Taylor and Francis, London.
28
Dolphin C.T., Beckett D.J., Janmohamed A., Cullingford T.E., Smith R.L., Shephard
E.A. and Phillips I.R. (1998) The flavin-containing monooxygenase 2 gene (FMO2) of
humans, but not of other primates, encodes a truncated, nonfunctional protein. J Biol
Chem, 273, 30599-30607.
Falls G.J., Blake B.L., Cao Y., Levi P.E., Hodgson E. (1995). Gender difference in
hepatic expression of flavin-containing monooxygenase isoforms (FMO1, FMO3 and
FMO5) in mice. J Biochem Toxicol, 10, 171-177.
Furnes B., Feng J., Sommer S.S. and Schlenk D. (2003). Identification of novel variants
of the flavin-containing monooxygenase gene family in African Americans. Drug
Metab Dispos, 31, 187–193.
Genter M.B., Deamer N.J., Blake B.L., Wesley D.S. and Levi P.E. (1995). Olfactory
toxicity of methimazole: dose-response and structure-activity studies and
characterization of flavin-containing monooxygenase activity in the Long-Evans rat
olfactory mucosa. Toxicol Pathol, 23, 477-486.
Haining R.L., Hunter A.P., Sadeque A.J., Philpot R.M. and Rettie A.E. (1997)
Baculovirus-mediated expression and purification of human FMO3: catalytic,
immunochemical, and structural characterization. Drug Metab Dispos, 25, 790-797.
Hardwick S.J., Skamarauskas J.T., Smith L. L., Upshall D.G. and Cohen G.M. (1991).
Protection of rats against the effects of alpha-naphthylthioruea (ANTU) by elevation of
non-protein sulphydryl levels. Biochem Pharmacol, 42, 1203-1208.
Hendersen M.C., Krueger S.K., Siddens L.K., Stevens J.F. and Williams D.E. (2004a).
S-oxygenation of the thioether organophosphate insecticides phorate and disulfoton by
human lung flavin-containing monooxygenase 2. Biochem Pharmacol, 68, 959-967.
Hendersen M.C., Krueger S.K., Stevens J.F. and Williams D.E. (2004b). Human flavincontaining monooxygenase form 2 S-oxygenation: sulfenic acid formation from
thioureas and oxidation of glutathione. Chem Res Toxicol, 17, 633-640.
Hernandez D., Janmohamed A., Chandan P., Philips I.R. and Shephard E.A. (2004).
Organization and evolution of the flavin-containing monooxygenase genes of human
and mouse; identification of novel gene and pseudogene clusters. Pharmacogenetics,
142, 117-30.
Hines R.N., Cashman J.R., Philpot R.M., Williams D.E. and Ziegler D.M. (1994). The
mammalian flavin-containing monooxygenases: molecular characterization and
regulation of expression. Toxicol Appl Pharmacol, 125, 1-6.
Hvattum E., Bergseth S., Pedersen C.N., Bremer J., Aarsland A. And Berge R.K.
(1991). Microsomal oxidation of dodecylthioacetic acid (a 3-thia fatty acid) in rat liver.
Biochem Pharmacol, 41, 945-953.
29
Janmohamed A., Dolphin C.T., Phillips I.R. and Shepard E.A. (2001). Quantification
and celluar localization of expression in human skin of genes encoding flavincontaining monooxygenases and cytochromes P450. Biochem Pharmacol, 62, 777-86.
Janmohamed A., Hernandez D., Phillips I.R. and Shephard E.A. (2004). Cell-, tissue-,
sex- and developmental stage-specific expression of mouse flavin-containing
monooxygenases (FMOs). Biochem Pharmacol, 68, 73-83.
Kaderick R.K., Wester E. and Ziegler M. (1991). Selective loss of liver flavincontaining monooxygenase in rats on chemically defined diets. Clin Pharmacol, 8, 95103.
Koukouritaki S.B., Simpson P., Yeung C.K., Rettie A.I. and Hines R.N. (2002). Human
hepatic flavin-containing monooxygenases 1 (FMO1) and 3 (FMO3) developmental
expression. Pediatr Res, 51, 236-234.
Krueger S.K., Siddens L.K., Martin S.R., Yu Z., Pereira C.B. (2004). Differences in
FMO2*1 allelic frequency between Hispanics of Puerto Rican and Mexican descent.
Drug Metab Dispos, 32, 1337-1340.
Krueger S.K. and Williams D.E. (2005). Mammalian flavin-containing
monooxygenases: structure/function, genetic polymorphisms and role in drug
metabolism. Pharmacol Therap, 106, 357-387.
Krueger S.K., Williams D.E., Yueh M.-F., Martin S.R., Hines R.N., Raucy J.L.,
Dolphin C.T., Shephard E.A. and Phillips I.R. (2002). Genetic polymorphisms of
flavin-containing monooxygenase. Drug Metab Rev, 34, 523-532.
Larsen-Su S., Krueger S.K., Yueh M.F., Lee M.Y., Shehin S.E., Hines R.N. and
Williams D.E. (1998). Flavin-containing monooxygenase isoform 2: developmental
expression in fetal and neonatal rabbit lung. J Biochem Mol Toxicol, 13, 187-193.
Larsen-Su S. and Williams D.E. (1996). Dietary indole-3-carbinol inhibits FMO
activity and the expression of flavin-containing monooxygenase form 1 in rat liver and
intestine. Drug Metab Dispos, 24, 927-931.
Lawton M.P., Cashman J.R., Cresteil T., Dolphin C.T., Elfarra A.A., Hines R.N.,
Hodgson E., Kimura T., Ozols J. and Phillips I.R. (1994). A nomenclature for the
mammalian flavin-containing monoxygenase gene family based on amino acid
sequence identities. Arch Biochem Biophys, 308, 254-257.
Lawton M.P., Gasser R., Tynes R.E., Hodgson E. and Philpot R.M. (1990). The flavincontaining monooxygenase enzymes expressed in rabbit liver and lung are products of
related but distinctly different genes. J Biol Chem, 265, 5855-5861.
30
Lawton M.P. and Philpot R.M. (1993). Functional characterization of flavin-containing
monooxygenase 1B1 expressed in Saccharomyces cerevisiae and Escherichia coli and
analysis of proposed FAD- and membrane-binding domains. J Biol Chem, 268, 57285734.
Lee P.W., Arnau T., and Neal R.A. (1980). Metabolism of alpha-naphthylthiourea by
rat liver and rat lung microsomes. Toxicol Appl Pharmacol. 53, 164-173.
Lomri N., Yang Z. and Cashman J.R. (1993a). Expression in Escherichia coli of the
flavin-containing monooxygenase D (form II) form adult human liver: determination of
a distinct tertiary amine substrate specificity. Chem Res Toxicol, 6, 425-429.
Lomri N., Yang Z. and Cashman J.R. (1993b). Regio- and stereoselective
oxygenations by adult human liver flavin-containing monoxygenase 3. Comparison
with Forms 1 and 2. Chem Res Toxicol, 6, 800-807.
McCombie R.R., Dolphin C.T., Povey S., Philips I.R. and Shepard E.A. (1996).
Localization of human flavin-containing monooxygenase genes FMO2 and FMO5 to
chromosomes 1. Genomics, 34, 426-429.
Myers C.R., Porgilsson B. and Myers J.M. (1997). Antibodies to a synthetic peptide
that react with flavin-containing monooxygenase (HLFMO3) in human hepatic
microsomes. J Pharmacol Toxicol Methods, 37, 61-66.
Nikbakht K.N., Lawton M.P. and Philpot R.M. (1992). Guinea pig or rabbit lung flavincontaining monooxygenases with distinct mobilities in SDS-PAGE are allelic variants
that differ in primary structure at only two positions. Pharmacogenetics, 2, 207-216.
Nnane I.P, and Damani L.A. (2001). Pharmacokinetics of trimethylamine in rats,
including the effects of a synthetic diet. Xenobiotica, 31, 749-755.
Overby L.H., Buckpitt A.R., Lawton M.P., Atta-Asafo-Adjei E., Schulze J. and Philpot
R.M. (1995). Characterization of flavin-containing monooxygenase 5 (FMO5) cloned
from human and guinea pig: evidence that the unique catalytic properties of FMO5 are
not confined to the rabbit ortholog. Arch Biochem Biophys, 317, 275-84.
Overby L.H., Carver G.C. and Philpot R.M. (1997). Quantitation and kinetic properties
of hepatic microsomal and recombinant flavin-containing monooxygenases 3 and
5 from humans. Chem Biol Interact, 106, 29-45.
Rouer E., Lemoine A., Cresteil T., Rouet P. and Leroux J.P. Effects of genetic or
chemically induced diabetes on imiramine metabolism. Drug Metab Disp, 15, 524-528.
Scott A.M., Powell G.M., Upshall D.G. and Curtis C.G. (1990). Pulmonary toxicity of
thioureas in the rat. Environ Health Perspect, 85, 43-50.
31
Shehin-Johnson S., Williams D.E., Larsen-Su S., Stresser D.M. and Hines R.N. (1994).
Tissue-specific expression of flavin-containing monooxygenase (FMO) forms 1 and 2
in the rabbit. J Pharmacol Exp Ther, 272, 1293-1299.
Shepard E.A., Dolphin C.T., Fox M.E., Povey S., Smith R. and Phillips I.R. (1993).
Localization of genes encoding three distinct flavin-containing monooxygenase to
human chromosome 1. Genomics, 16, 85-89.
Taylor K.H. and Ziegler D.M. (1987). Studies on substrates of the hog liver flavincontaining monooxygenase: anionic organic sulfur compounds. Biochem Pharmacol,
36, 141-146.
Whetstine J.R., Yueh M.F., McCarver D.G., Williams D.E., Park C.S., Kang J.H., Cha
Y.N., Dolphin C.T., Shephard E.A., Phillips I.R. and Hines R.N. (2000) Ethnic
differences in human flavin-containing monooxygenase 2 (FMO2) polymorphisms:
detection of expressed protein in African-Americans. Toxicol Appl Pharmacol, 168,
216-224.
Yeung C.K., Lang D.H., Thummel K.E. and Rettie A.E. (2000). Immunoquantitation of
FMO1 in human liver, kidney, and intestines. Drug Metab Dispos, 28, 1107-1111.
Yueh, M.-F., Krueger S.K. and Williams D.E. (1997). Pulmonary flavin-containing
monooxygenase (FMO) in rhesus macaque: expression of FMO2 protein, mRNA and
analysis of the cDNA. Biochim Biophys Acta, 1350, 267-271.
Ziegler D.M. (1984). Metabolic oxygenation of organic nitrogen and sulfur compounds.
Drug Metabol Drug Tox. (J.R. Mitchell and M.G. Horning eds), 33-53. Raven Press,
New York.
Ziegler D.M. (1991). Bioactivation of xenobiotics by flavin-containing
monooxygenases. Adv Exp Med Biol, 283, 41-50.
Ziegler D.M. and Mitchell C.H. (1972). Microsomal oxidase IV: properties of mixedfunction amine oxidase isolated from pig liver microsomes. Arch Biochem Biophys,
150, 116-25.
32
APPENDIX
(a)
Mass of Female Mice on NIH31 Diet
20
18
16
14
12
10
M ouse 1
8
M ouse 2
6
M ouse 3
4
M ouse 4
2
0
1
3
5
7
9
11
13 15 17 19
Time (Days)
21
23
25
27
29
31
(b)
Mass of Female Mice on AIN93G Diet
20
18
16
14
12
10
M ouse 1
8
M ouse 2
6
M ouse 3
4
M ouse 4
2
0
1
3
5
7
9
11 13 15 17 19 21 23 25 27
Time (Days)
29 31
33
(c)
Mass of Male Mice on NIH31 Diet
22
20
18
16
14
12
10
M ouse 1
8
M ouse 2
6
M ouse 3
4
M ouse 4
2
0
1
3
5
7
9
11
13
15 17 19
Time (Days)
21
23
25
27
29
31
(d)
Mass of Male Mice on AIN93G
20
18
16
14
12
10
M ouse 1
8
M ouse 2
6
M ouse 3
4
M ouse 4
2
0
1
3
5
7
9
11
13 15 17 19
Time (Days)
21
23
25
27
29
31
Figure A1. Mass of 129 S/V mice from 3-5 weeks of age. (a) Female on NIH31 diet,
(b) Female on AIN93G diet, (c) Male on NIH31 diet and (d) Male on AIN93G diet
34
Figure A2. Agilent 2100 Bioanalyzer Electropherogram, representative of mouse lung
total RNA demonstrating acceptable quality and integrity
35
Table A1. Composition of NIH31 diet
Ingredients (%)
Crude oil
Crude protein
Crude fiber
Ash
Nitrogen free extract
Dietary Fiber
Starch
Sugars
Minerals
Calcium
Phosphorus
Sodium
Chloride
Potassium
Magnesium
Iron
Manganese
Zinc
Copper
Iodine
Cobalt
Selenium
Chromium
%
%
%
%
%
%
%
%
NIH31
4.30
19.00
4.00
6.10
53.70
12.80
38.00
6.30
%
%
%
%
%
%
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
mg/kg
1.00
0.65
0.29
0.54
0.93
0.22
185.00
118.00
67.00
13.00
11.00
0.50
0.18
0.50
36
Table A2. Composition of AIN93G diet
Ingredients (%)
Casein
Dextrinized cornstarch
Sucrose
Corn starch
Alphacel, Non-nutritive bulk
Soybean oil
AIN-93M mineral mix*
L-Cystine
AIN-93M-VX vitamin mix
Choline Bitartrate
tert-Butylhydroquinone
Minerals
Calcium carbonate
Monopotassium phosphate
Potassium citrate monohydrate
Sodium chloride
Potassium sulfate
Magnesium oxide
Ferric citrate
Zinc carbonate
Manganese carbonate
Copper carbonate
Potassium iodate
Sodium selenate, anhydrous
Ammonium Molybdate 4H2O
Sodium Metasilicate 9H2O
Chromium Potassium Sulfate 12 H2O
Lithium chloride
Boric acid
Sodium fluoride
Nickel Carbonate
Ammonium Vanadate
Powdered sugar
*Contains ingredients listed in Minerals
%
%
%
%
%
%
%
%
%
%
%
AIN93G
14.0000
15.5000
10.0000
46.6000
5.0000
4.0000
3.5000
0.1800
1.0000
0.2500
0.0008
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
%
35.7000
19.6000
7.0780
7.4000
4.6600
2.4000
0.6060
0.1650
0.0630
0.0300
0.0010
0.0010
0.0008
0.1450
0.0275
0.0017
0.0081
0.0064
0.0032
0.0007
22.1000
37