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Department of Nutritional Sciences, University of Arizona, Tucson, AZ 85721
3To whom correspondence and reprint requests should be addressed at Department of Nutrition and Food Science, University of Maryland, College Park, MD 20742.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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KEY WORDS: • hamsters • dietary zinc • cardiovascular disease • cholesterol • low density lipoprotein
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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,
Powell et al. 1987
) known as apoB mRNA editing. A
cytidine to uridine conversion in the apoB transcript results in
changing a glutamine CAA codon to UAA, an in-frame translational
stop codon. This editing results in a truncated protein (apoB-48) that
is approximately half the size of apoB-100. ApoB-48 is identical to the
amino-terminal half of apoB-100 but lacks the COOH-half of
apoB-100. Consequently, apoB-48 lacks the LDL receptor-binding
domain required for uptake via LDL receptor–mediated endocytosis.
Instead, apoB-48–containing lipoproteins are cleared faster than
apoB-100–containing lipoproteins by a distinct pathway. ApoB-48 also
does not possess the region present in apoB-100 that can combine with
apo(a) to form the atherogenic lipoprotein (a) molecule (Scanu and Fless 1990
). Because humans do not have the ability to edit
apoB mRNA in the liver, human VLDL and LDL particles contain apoB-100.
This lack of hepatic editing activity is thought to contribute to our
susceptibility to develop atherosclerosis. Consequently, numerous
laboratories are developing ways to induce hepatic editing using
techniques such as gene therapy.
The editing reaction occurs in the nucleus (Lau et al. 1991
) and is performed by a complex or "editosome"
(Harris et al. 1993
) comprised of a 27-kDa enzyme termed
apoB mRNA editing catalytic subunit
1 (apobec-1) and other complementary protein factor(s)
(Lau et al. 1990
, Navaratnam et al. 1993b
). Apobec-1 exists as a homodimer (Lau et al. 1994
) and is an RNA-specific cytidine deaminase that
exhibits considerable sequence homology with other cytidine deaminases
(Navaratnam et al. 1995
). A zinc-coordinating region
that resembles that of other cytidine deaminases is present in
apobec-1, and apoB mRNA editing has been shown to be zinc dependent in
vitro (Barnes and Smith 1993
, Navaratnam et al. 1993a
). Editosome assembly is facilitated by an 11-nucleotide
"mooring" sequence that begins five nucleotides downstream of the
editing site in the apoB transcript (Smith et al. 1991
);
this region has been shown to be critical for editing efficiency
(Shah et al. 1991
). Although the complementary factors
are absolutely essential for editing (Anant et al. 1995
,
Teng et al. 1993
), they are present in a wide variety of
tissues including some that do not edit apoB mRNA (Teng and Davidson 1992
). Because of the ubiquitous nature of the
complementary factors, it often appears to be the tissue-specific
expression of apobec-1 that determines the ability or inability of a
tissue to edit. Although intestinal editing occurs at high levels in
nearly all mammals examined, hepatic editing is much less common and
occurs only in certain species. Greeve et al. (1993)
compared hepatic editing in 12 different species and showed that mice,
rats, dogs and horses exhibit hepatic editing. Humans, pigs, cows,
monkeys, sheep and hamsters, as we have reported here, do not possess
detectable hepatic editing. This finding may explain in part why
hamster lipoprotein metabolism is more similar to humans than that of
most other rodents. For example, a relatively high portion of plasma
cholesterol is LDL cholesterol in both humans and hamsters. Both
species respond similarly to atherogenic diets (Spady and Dietschy 1988
), and hamsters also develop atherosclerotic
lesions that resemble those found in humans (Nistor et al. 1987
). These characteristics make the hamster an intriguing
model with which to study lipoprotein metabolism and prompted us to
examine apoB mRNA editing in hamsters.
Previous work in other laboratories has shown that hamster hepatocytes
secrete apoB-100 but not apoB-48 (Liu et al. 1991
).
These studies did not define the mechanism for the lack of apoB-48
synthesis. We hypothesized that hamsters do not synthesize apoB-48 in
the liver because they do not edit apoB mRNA in the liver due to a lack
of expression of apobec-1, the catalytic component of the editing
reaction. Because the editing reaction is zinc dependent in vitro and
zinc-deficient rats exhibit impaired lipid absorption, we
hypothesized that dietary zinc status may affect intestinal editing
activity in hamsters. Similarities between hamster and human
lipoprotein metabolism and the prevalence of zinc deficiency in certain
human subpopulations (Sandstead 1995
) prompted us to
examine whether zinc deficiency in hamsters affected intestinal editing
levels.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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For hepatic RNA isolation, fresh liver sample was immediately combined
with TRIzol reagent (Life Technologies, Grand Island, NY) and
homogenized. Subsequent steps were performed according to the
manufacturer’s protocol. For intestinal RNA isolation, the small
intestine was flushed with ice-cold diethyl pyrocarbonate
(DEPC)-treated PBS and the mucosa was obtained by scraping. The
intestinal mucosa was immediately combined with TRIzol for
homogenization and isolation of intestinal RNA. Other tissue samples
were snap-frozen in liquid nitrogen immediately after their
excision and stored at -80°C for subsequent RNA isolation. RNA
samples were digested with RQ1 DNase in a 60-µL
reaction containing 
5 µg RNA, 5 µL
RQ1 DNase (1 U/µL; Promega, Madison, WI),
1 µL RNasin (40 U/µL;
Promega), 6 µL of 100 mmol/L dithiothreitol (DTT), 6
µL of 10X RQ1 buffer and 37 µL
DEPC-H2O. Samples were digested for 90 min at 37°C. RNA
was then extracted with phenol/chloroform, precipitated with ethanol
and pellets were resolubilized in 10 µL
DEPC-H2O.
Cloning of apoB cDNA sequence flanking the editing site.
Human and rat apoB cDNA sequences flanking the editing site were obtained from GenBank (NIH, Bethesda, MD), and the conserved regions were analyzed by Genetics Computer Group (GCG) program (Madison, WI). A pair of primers, RP1–5' and RP1–3', were then designed on the basis of the rat sequence. DNase I-digested hamster hepatic or intestinal RNA (500 ng) were mixed with 2 µL of 5X first-strand synthesis buffer (Life Technologies), 50 pmol RP1–3' primer and DEPC-water to 10 µL total volume. The sample was heated to 70°C for 10 min, then cooled to 37°C for 15 min, during which time 4 µL reverse transcription (RT) buffer, 2 µL of 10 mmol/L dNTPs and 1 µL of MMLV RT (Life Technologies) were added. Reverse transcription was performed at 42°C for 30 min, then stopped by heating to 95°C for 5 min and brief placement on ice. Thirty cycles of polymerase chain reaction (PCR) were then performed with 50 pmol RP1–5' primer, and the cycling condition was set to 92°C for 1 min, 42°C for 2 min and 65°C for 2 min. The resultant PCR products were cloned, and the positive clones were verified by sequence analysis. Multiple clones were derived and sequenced from separate PCR reactions to reduce the possibility of PCR-introduced mutations. The editing site for hamster apoB was deduced from the sequence alignment of hamster, human, rat, pig and mouse cDNA flanking region of the editing site, and was further confirmed by the comparison of apoB sequences derived from both hamster liver and intestine. Oligo sequences were as follows: 5'-TCCTCAGCAGATTCATGATTATCT-3' (RP1–5'; rat apoB cDNA nt 6482–6505), 5'-AGCATTTTTAGCTTTTCAATGATT-3' (RP1–3'; rat apoB cDNA nt 6740–6763).
Cloning of the hamster apobec-1 cDNA.
The hamster apobec-1 cDNA was obtained by the method of RT-PCR, as
described previously (Wu et al. 1998
) with some
modifications. Apobec-1 mRNA sequences for humans, rats and mice were
obtained from the GenBank database and analyzed for conserved regions.
Three primers, Fa, Ra and Rb, were then designed on the basis of the
human sequence. RT was performed with DNase I–digested hamster
intestinal RNA, primer Ra and rTth (Perkin-Elmer, Norwalk, CT). The
mixture was heated at 70°C for 10 min, annealed at 55°C for 10 min
and the reverse transcription performed at 70°C for 10 min. For PCR,
8 µL of 10X chelating buffer, 50 pmol Fa primer, 6
µL of 25 mmol/L MgCl2, and 64
µL distilled water were added. After 3 min at 94°C,
PCR was performed for 30 cycles in a Perkin-Elmer thermocycler as
follows: 94°C/1 min, 55°C/2 min and 72°C/2 min, with a 10-min
final extension at 72°C. To reduce downstream selection and
verification workload, the resultant PCR fraction was then reamplified
with primers Fa and Rb, and this PCR resulted in a 500-bp product.
This fragment was cloned and found to share high homology to all known
apobec-1 mRNA sequences; it was designated as the central part of
hamster apobec-1 cDNA. On the basis of this sequence, two hamster
primers, Fb and Rc, were designed for cloning the 5' and 3' ends of
hamster apobec-1 cDNA using the method of RACE (Rapid Amplication of
cDNA Ends).
For 5' RACE, the RT reaction was performed as described above with the
primer Rb; samples were prepared as previously described (Wu et al. 1998
). To enhance the selectivity, the secondary PCR with
primers dT18-RI and Rc was then performed. The resultant PCR product
was cloned and verified by sequencing analysis. In 3' RACE, the RT
reaction was first performed with dT18-RI primer, using MMLV, and then
amplified with dT18-RI and primer Fa. The resultant PCR was divided
into four aliquots, and further amplified with the primers Fb and one
of dT16GACG, dT16GACA, dT16GACT or dT16GACC. Positive bands were
observed in the reactions from dT16GACA and dT16GACC. The PCR products
were cloned and verified by sequencing analysis. Multiple clones were
derived from separate PCR reactions to reduce the likelihood of
sequence mutations during PCR. All positive clones derived from 5' and
3' RACE were analyzed with GenBank sequence homology search, and the
one containing the longest insert was selected as hamster apobec-1 5'
or 3'-end cDNA.
Sequences for the 5', central, and 3' portions of apobec-1 cDNA were analyzed by GCG program to compile the full-length cDNA sequence. Primers used to obtain the hamster apobec-1 cDNA sequence were as follows: 5'-cggaattctttttttttttttttttt-3' (dT18-RI), 5'-gtgaccccactctgaggagaagaat-3' (Fa; corresponds to hamster apobec-1 cDNA nt 76–99), 5'-TTCCTGTCCTGGAGTCCCTG-3' (Fb; hamster apobec-1 cDNA nt 305–324), 5'-tcttcttgaaatctttaaacagggtgg-3' (Ra; corresponds to hamster apobec-1 cDNA nt 614–640), 5'-tacagcctcatccacagattt-3' (Rb; corresponds to hamster apobec-1 cDNA nt 559–579), 5'-ATTTGAAGGTGGGTAGTTGAC-3' (Rc; hamster apobec-1 cDNA nt 515–535). Upper-case letters denote hamster apobec-1 cDNA sequences.
ApoB mRNA editing assay.
ApoB mRNA editing was determined by RT-PCR amplified primer
extension assay, basically as described by Giannoni et al. (1994)
. This increases the sensitivity of the assay but does
not change the ratio of apoB-48/apoB-100 mRNA. dNTPs (1 mmol/L), 4
µL of 5X first-strand synthesis buffer (Life
Technologies), 1 µL RNasin (40
U/µL; Promega), 2 µL of
100 mmol/L DTT, and 50 pmol of downstream primer RP1–3' were combined
with 8 µL DNase-digested RNA (750 ng) in a
19-µL reaction. Samples were heated to 70°C for 7
min, then annealed at 42°C for 5 min. SuperScript II RT (40
U; Life Technologies) was added and samples were incubated
for 60 min at 37°C. After reverse transcription, mineral oil was
added; samples were heated to 95°C for 5 min and then placed briefly
on ice. For PCR, 80 µL of PCR buffer that contained 10
µL of 10X PCR buffer (Perkin-Elmer), 2
µL (50 pmol) primer RP1–5', 10 µL
dimethyl sulfoxide (DMSO), 10 µL of 25 mmol/L
MgCl2, 1 µL Taq polymerase (Perkin-Elmer)
and 47 µL distilled H2O were added to the
20-µL reaction followed by 30 cycles of 92°C/1 min,
40°C/2 min and 65°C/2 min. For the last cycle, extension was
performed for 10 min. PCR with these primers resulted in the
amplification of a 282-bp fragment flanking the editing site. PCR
products were purified with QIAquick PCR purification kit (Qiagen,
Valencia, CA) after size verification by agarose gel electrophoresis.
To check for DNA contamination, a control for each RNA sample was kept
on ice during RT; in addition, PCR was performed using Taq DNA
polymerase (Perkin-Elmer). Both methods verified the absence of DNA
contamination. For primer extension analysis, the annealing primer
HAMBEDIT (spanning hamster apoB cDNA nt 6674 to 6708) was 5'-end
labeled with (
-32P)-ATP (Amersham), then purified with a
QIAquick Nucleotide Removal Kit (Qiagen). Labeled primer, 2
µL of 250 mmol/L NaCl and 2 µL of 5X
first-strand synthesis buffer (Life Technologies) were combined
with 1 µL of diluted purified PCR product. Tubes were
heated at 95°C for 10 min and after annealing at 70°C for 2 min,
the mixture was cooled to 37°C over 2 min. Thereafter, 20
U of SuperScript II RT (Life Technologies), 1
µL of 100 mmol/L DTT, 1 µL of 5
mmol/L ddGTP and 0.5 µL of a 10 mmol/L mixture of
dATP, dCTP and dTTP were added. Extension was performed at 37°C for
30 min. Samples were then placed on ice, and the products were
ethanol-precipitated and separated on 6% polyacrylamide/8 mol/L
urea gels at 55 W for 2 h. Gels were dried, subjected to
autoradiography and analyzed by laser densitometry (Molecular Dynamics,
Sunnyvale, CA). Primer extension of the unedited apoB-100 mRNA resulted
in a product 43 bases in length, whereas the edited apoB-48 mRNA
product was 54 bases in length. Primers used for the editing assay,
5'-AATCCTGTGGATCATAATTGTCTCTAATATACTGA-3' (HAMBEDIT), RP1–3' and
RP1–5', were described in the section on cloning of the apoB cDNA
sequence flanking the editing site.
Tissue-specific distribution of apoB and apobec-1 mRNA.
RT-PCR was used to amplify endogenous mRNA of both apobec-1 and apoB in
the various tissues. Competitive RT-PCR was not utilized because a
semiquantitative method was not required. For apoB, RT was performed as
previously described in the last section. RT product (10
µL) was combined with 40 µL PCR mix
in a 50-µL final volume containing 50 pmol RP1–5', 5
µL of 10X PCR buffer (Perkin-Elmer), 2
µL of 25 mmol/L MgCl2, 5
µL DMSO, 0.5 µL Taq polymerase and
1.5 µL diluted [
-32P] dCTP (NEN Life
Science Products, Boston, MA). The first cycle was performed as
follows: 92°C/3 min, 40°C/2 min and 65°C/2 min. Twenty-nine
cycles of 94°C/1 min, 55°C/2 min and 72°C/2 min were then
performed, followed by a 10-min final extension.
For apobec-1, RT was performed essentially as described previously
except that 50 pmol Rc primer was used in a 10-µL RT
reaction. For subsequent PCR, 40 µL of PCR buffer
containing 4 µL of 10X chelating buffer, 50 pmol Fb, 3
µL of 25 mmol/L MgCl2 and 0.15
µL [-32P] dCTP were added. After 2
min at 94°C, 30 cycles were performed as follows: 94°C/30 s,
55°C/30 s and 72°C/2 min, with a 10 min final extension. Negative
controls for all samples were included in which tubes were kept on ice
during reverse transcription (RT-) to check for DNA contamination.
RT-PCR products were electrophoresed on native 5% polyacrylamide
gels. Migration of the products was compared with a 100-bp DNA ladder
(Promega) 5' end-labeled with [-32P]
ATP.
Experimental animals and diets.
Male Golden Syrian hamsters (7 wk old; Harlan, Indianapolis, IN) were
assigned randomly to one of the following three dietary treatment
groups: zinc-adequate (ZA, n = 8),
zinc-deficient (ZD, n = 8) or
zinc-replenished (ZDA, n = 4). The replenished
group was fed ZD diet until the last 2 d of treatment during which
time they were fed the ZA diet. Dietary treatment lasted for 49 d
during which time hamsters consumed their respective diet ad libitum.
The basal diet was purchased from Dyets (Bethlehem, PA) and was
formulated according to the AIN-93-M rodent diet recommendations
(Reeves et al. 1993
) except that zinc was omitted from
the mineral mix. ZnCO3 was added to the zinc-adequate
diet at 30 mg Zn/kg diet, whereas the zinc-deficient diet contained
<0.5 mg Zn/kg diet. All procedures were approved by the Animal Care
and Use Committee of the University of Arizona.
Statistics.
Values are means ± SD. Differences were considered
significant at P < 0.05. The data were analyzed
using one-way ANOVA and Duncan’s new multiple range test
(Jaccard and Becker 1990
).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The hamster apobec-1 cDNA was cloned by RT-PCR and 5', 3'-RACE. The
complete nucleotide and the predicted amino acid sequence are shown in
Figure 1
. The nucleotide sequence is 829 bp in length and contains a consensus
polyadenylation signal (unlike the rat in which the consensus signal is
missing). The single open reading frame of 690 bp encodes a 229-amino
acid protein with a calculated molecular mass of 27,574 Da. As
indicated in Figure 1
, the hamster, mouse and rat proteins each contain
229 residues, whereas those of humans and rabbits are each 236 residues
in length. The highest similarity is with mouse apobec-1, sharing 83%
amino acid sequence identity, followed by rat (81%), human (69%) and
rabbit (64%).
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A 281-bp hamster apoB cDNA fragment was obtained by RT-PCR. The
editing site is identified by sequence alignment with known apoB
sequences, as shown in Figure 2
. Only one nucleotide was found to be different when comparing the apoB
sequences derived from hamster liver and intestine, at the editing
site, indicating that the PCR method was accurate and did not introduce
mutations. Alignment of the apoB mRNA sequences once again illustrates
the importance of the sequence downstream of the editing site that has
been shown to be critical for editing (Shah et al. 1991
). Smith et al. (1991)
proposed that
nucleotides 6671–6681, 3' to the editing site (nt 6666), constitute a
"mooring sequence" in which the sequence functions to
position the deaminase over the site to be edited. Indeed, a span of 18
nucleotides downstream of the editing site is conserved in each of
the species shown in Figure 2
.
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Using the sequence derived from the apoB mRNA adjacent to the editing
site, we synthesized an annealing primer for the "poisoned" primer
extension assay used to measure editing. RNA was prepared from 12
different tissues, the region of apoB mRNA flanking the editing site
was amplified by PCR; then, the primer extension editing assay was
performed. The presence of ddGTP in the reaction causes termination of
extension when a CTP is in the template strand, the template being apoB
mRNA, thereby allowing differentiation between edited (apoB-48) and
unedited (apoB-100) transcripts. Results from the editing assays are
shown in Figure 3
. We were not surprised to find the highest level of editing in the
small intestine and colon RNA samples. However, there also appeared to
be very small amounts of editing in the stomach and pancreas because an
apoB-48 product was barely visible. Of particular interest is the
finding that hamsters apparently do not edit apoB mRNA in the liver or
at least the editing is at an extremely low level, below detection by
primer extension analysis. To the best of our knowledge, this
represents the first published report using primer extension analysis
to establish that hamsters do not edit apoB mRNA in the liver. These
findings support those of other researchers indicating that hamsters do
not synthesize apoB-48 protein in the liver.
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Because the editing assay detected low levels of editing activity in
several tissues, we chose to examine the tissue-specific expression
of apobec-1. Initially, we had set up ribonuclease protection assays
(RPA) to detect apoB and apobec-1 mRNA in the various tissues, using
the newly cloned cDNA fragments as probes. Low levels of apobec-1 mRNA
expression were detected in the intestine but not in any other tissues
(data not shown). In contrast, apoB mRNA was detected in both liver and
intestine, at relatively high expression levels, but no clear RPA bands
for other tissues were observed (data not shown). Because the editing
assay detected editing activity in tissues other than the small
intestine, we then selected the more sensitive RT-PCR method to
search for apoB and apobec-1 expression. For apobec-1, the most
important RT-PCR product was found in the small intestine; however,
much lower levels of apobec-1 mRNA were also present in other tissues,
including a very slight but visible band in hamster liver RNA
(Fig. 4
). RT-PCR of apoB mRNA showed that the highest level of apoB mRNA
was present in the liver and small intestine (Fig. 5
). Slight bands were also present in the colon, kidney, stomach and
pancreas samples (Fig. 5)
. Because the purpose of RT-PCR was to
examine tissue distribution and not to quantify apoB and apobec-1
mRNA abundance, we did not utilize a method such as competitive
RT-PCR for these studies.
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As mentioned earlier, the editing reaction has been shown to be a
zinc-dependent reaction in vitro (Barnes and Smith 1993
, Navaratnam et al. 1993a
). We reported
previously that rats consuming a zinc-deficient diet exhibited
modest decreases in plasma apoB-48 and hepatic editing levels
(Reaves et al. 1999
). To examine zinc dependency in
vivo, we fed hamsters a zinc-deficient diet for 7 wk. Because no
hepatic editing activity was detected, only intestinal editing was
measured. Hamsters consuming the zinc-deficient diet exhibited
signs of zinc deficiency such as depressed food intake, reduced growth
rate, lowered plasma zinc and reduced hepatic zinc. These findings were
reported previously in a study that examined the effect of zinc status
on apolipoprotein A-I gene expression (Wu et al. 1998
). Interestingly, the zinc-deficient hamsters exhibited
a reduction in intestinal apoB mRNA editing (Fig. 6
). Although the reductions were not drastic, they were significant.
Zinc-deficient hamsters that consumed a zinc-adequate diet for
the last 2 d of treatment (ZDA group) had higher editing levels
than ZD hamsters and were not different from ZA hamsters (Fig. 6)
. The
food intake of ZDA hamsters was not different from that of ZD hamsters
during the 2-d replenishment, remaining significantly lower than that
of ZA hamsters (Wu et al. 1998
).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). Experiments
using point mutations of these residues (Navaratnam et al. 1995
, Yamanaka et al. 1994
) have substantiated
the relevance and essentiality of these amino acids. The
leucine-rich region of the carboxyl-terminus, thought to play a
role in dimerization, is also relatively well conserved. The region
that spans residues 173–210 is highly conserved in hamsters, mice,
rats and humans. In fact, each leucine residue is conserved in these
four species at every instance except residue 196. At this position,
both hamsters and humans have residues other than leucine. It should
also be noted that as in the other species, the hamster sequence also
contains proline residues, suggesting that this region more closely
resembles that of a leucine-rich motif, previously shown to
facilitate protein-protein interactions (Roth 1991
),
instead of a classic leucine-zipper motif. Although the precise
function of this region is yet to be defined clearly, the importance of
its presence appears to be validated by its conservation across
species.
For the cloned hamster apoB cDNA fragment, the most interesting region
is just downstream of the editing site proposed to contain the
"mooring sequence" (Smith et al. 1991
). Indeed, the
region is perfectly conserved in each of the species shown in Figure 2
,
thereby lending further credence to its importance. In fact, when
considering the sequences in Figure 2
as well as those reported by
Greeve et al. (1993)
, out of a total of 11 different
species, only the guinea pig contains a one-nucleotide deviation in
the sequence spanning nucleotides 6666–6681. The significance of this
single nucleotide difference seems minimal, however, because the guinea
pig has relatively high intestinal editing (87%).
For apoB mRNA editing to occur, apobec-1, the catalytic subunit,
complementary factor(s) and apoB mRNA must all be present. Most data
point to the presence or absence of apobec-1 as determining whether a
tissue edits apoB mRNA; however, there are exceptions. There are some
tissues that edit apoB mRNA but do not synthesize and secrete apoB-48
protein (Teng et al. 1990
). Obviously, some tissues
express apoB mRNA but do not edit; however, it is not clear whether
there are tissues that express apobec-1 and apoB mRNA but do not have
editing activity. In these studies, possibly the most difficult finding
to interpret was the apparent lack of editing activity in the liver and
kidney in which both apobec-1 and apoB mRNA were detected, albeit at
very low levels. One explanation could be that the level of editing
activity was extremely low and was below the level of detection for
this assay. In the liver, apobec-1 may be a limiting factor, whereas in
the kidney, apoB mRNA may be limiting. Other possibilities could be
that the sensitivity of RT-PCR allows for detection of apobec-1
mRNA at levels that would expectedly yield very low levels of protein.
Post-transcriptional mechanisms that inhibit production of apobec-1
protein could also be involved. Examples exist in which apoB mRNA has
been detected in several tissues without apoB protein. The sensitivity
of RT-PCR is illustrated by our RPA findings (data not shown) in
which apobec-1 mRNA was seen only in the small intestine. Apobec-1
detection will obviously differ depending on which method is used;
hence it is often difficult to compare the results from different
studies. For example, in humans, Hadjiagapiou et al. (1994)
detected apobec-1 mRNA in stomach and testis tissue in
addition to the intestine by using RT-PCR analysis. However,
Lau et al. (1994)
reported that apobec-1 mRNA was
expressed only in the small intestine of humans as indicated by
Northern blot analysis. Most other studies have not reported both
apobec-1 expression and editing activity. We felt it was important to
investigate this in hamsters.
We chose to use RT-PCR analysis because of its sensitivity, although this assay poses some difficulties that are not encountered in Northern blots. Because we were not intending to use this method to quantitate mRNA abundance, no internal reference or competitor was used during RT-PCR. In view of its reported sensitivity, we used rTth to amplify apobec-1 mRNA. Apparently the use of SuperScriptII or rTth can result in slightly different results in the RT- reactions. For example, in the RT- reactions of apoB mRNA using SuperScriptII, no band is evident. However, when the same RNA samples were used for RT-PCR of apobec-1 with rTth, slight bands were visible in some of the RT- reactions. Therefore, these bands are most likely related to rTth and could be from a "leaking" of reverse transcriptase activity through the incubation on ice or during PCR.
Apobec-1 and/or apoB mRNA was present in tissues that would apparently
not require apoB protein synthesis. Why apobec-1 and/or apoB mRNA would
be expressed in these tissues is unknown, but this phenomenon has also
been observed by other researchers in several different species
(Hadjiagapiou et al. 1994
, Teng et al. 1990 and 1993
).
Dietary treatments reported in this study have been used routinely in
our laboratory to induce zinc deficiency in hamsters. We found that
changes in plasma and hepatic zinc concentrations were comparable
between hamsters consuming a zinc-deficient diet for 7 wk and rats
consuming a zinc-deficient diet for 2.5 wk (Wu et al. 1998
). Impaired absorption of dietary lipids has been
established in ZD rats (Koo and Turk 1977
, Koo et al. 1986
). In the intestinal absorptive cells, nascent
chylomicrons appear to be irregular in shape and larger in the
zinc-deficient than in zinc-adequate rats (Koo et al. 1985
). Because of these findings, and the fact that editing has
been shown to be zinc dependent in vitro, we examined the influence of
dietary zinc on intestinal editing levels. Although hepatic editing has
been shown to be influenced by several nutritional regimens
(Baum et al. 1990
, Funahashi et al. 1995
,
Leighton et al. 1990
, Reaves et al. 1996 and 1999
), intestinal editing activity has been found to be
resistant to dietary manipulations. In the past, we reported that
copper deficiency in rats increased the plasma ratio of apoB-48 to
total apoB, as well as hepatic editing activity. However, possibly due
to high variances, the intestinal editing activity was not different
among the treatments (Reaves et al. 1996
). One
explanation for these observations is that high intestinal editing
activity is the result of highly saturated apobec-1 enzymatic activity.
As a result, small changes in the level of apobec-1 will not lead to
detectable changes in editing activity. In this study, however, our ZD
hamsters exhibited a small but significant reduction in intestinal
editing. Refeeding a ZA diet to ZD hamsters for the last 2 d
resulted in the restoration of intestinal editing activity to a level
comparable to that of ZA hamsters. The food intake for the ZDA group
was not increased during the 2-d repletion, as was reported previously
(Wu et al. 1998
). Our previous work in rats suggested
that differences in food consumption were not responsible for returning
ZDA hepatic editing to ZA levels (Reaves et al. 1999
).
In a previous publication, we reported the measurement of mRNA
abundance of the well-described zinc-responsive metallothionein
(MT)-II gene as a means of assessing zinc levels in the intestinal
mucosal cells. Tissue samples used for the determination of editing
activity in this study and those used for MT-II analysis were
derived from the same hamsters. The previous work indicated that
intestinal MT-II mRNA levels were reduced by 71% in ZD hamsters
compared with ZA hamsters. In the ZDA group, MT-II mRNA was
elevated to 259% of ZA values (Wu et al. 1998
). These
data strongly suggest that intestinal zinc levels were different among
groups and that zinc levels in the intestinal mucosal cells were
restored by the zinc replenishment. In view of the metallothionein data
as well as the food intake data, this would suggest that differences in
intestinal editing are due to differences in cellular zinc levels.
We did not report the hamster plasma apoB-48/apoB-100 ratios for several reasons. ApoB-48 in hamsters is derived from intestine. We observed very low levels of apoB-48 in hamster plasma (unpublished observation). These low levels of apoB-48 with the relatively high levels of apoB-100 make it difficult to quantify apoB-48/apoB-100 ratios in an accurate manner. We realize that there are several other aspects of apoB metabolism that might be affected by zinc deficiency. Alterations in synthesis, degradation, secretion and/or circulatory half-life of apoB-48 derived from the intestine could all be affected potentially by zinc status. However, we chose to focus our efforts on apoB editing.
It should be noted that our current findings differ from those of a
study by Nassir et al. (1996)
in which neither hepatic
nor intestinal editing levels were changed significantly by the
consumption of a zinc-deficient diet in rats. However, we reported
recently that rats consuming a zinc-deficient diet do indeed
exhibit reductions in hepatic editing levels compared with
zinc-adequate controls (Reaves et al. 1999
). As
pointed out in our previous study, there are many differences in the
experimental design between the two studies that could explain distinct
observations regarding hepatic editing. However, it may be extremely
difficult to compare findings from these two studies with the current
work on hamsters because of fundamental differences in lipoprotein
metabolism that set these species apart.
In summary, we demonstrated that hamsters have no or extremely low
levels of hepatic editing activity. We also reported the cDNA sequence
of hamster apobec-1 and apoB mRNA sequence flanking the editing site.
Moreover, our findings indicate that intestinal editing activity was
reduced in hamsters consuming a zinc-deficient diet. Although it
may not be common for humans to consume a diet as low in zinc as the
diets these studies, there are additional factors in humans that may
promote zinc deficiency. Dietary factors that have been shown to
promote zinc deficiency include diets high in phytates (O’Dell and Savage 1960
), certain types of fiber (Reinhold et al. 1976
) and casein (Harzer and Kauer 1982
). In
addition, other factors such as age (Prasad et al. 1993
), excessive alcohol use, liver diseases, malabsorption
syndromes, renal disease, enteral or parenteral alimentation, and
sickle cell disease (as reviewed by Evans 1986
) may also
increase the risk of zinc deficiency. A diet low in zinc and/or any
number of the above factors could contribute to compromised zinc status
in humans. Moreover, severe zinc deficiency has been observed in humans
with a rare, autosomal recessive disease termed acrodermatitis
enteropathica (Barnes and Moynahan 1973
). Therefore,
data derived from the zinc-deficient hamster model can provide
useful information concerning apolipoprotein B metabolism in humans.
|
FOOTNOTES |
|---|
2 The nucleotide sequences reported in this paper have been submitted to GenBank with accession numbers AF176576 (apoB) and AF176577 (apobec-1).
4 Abbreviations used: apoB, apolipoprotein B; apobec-1, apoB mRNA editing catalytic subunit 1; DEPC, diethyl
pyrocarbonate; DMSO, dimethyl sulfoxide; DTT, dithiothreitol; GCG, Genetics Computer Group; MT, metallothionein; RACE, Rapid Amplication of cDNA Ends; RPA, ribonuclease protection assay; RT-PCR. reverse transcriptase-polymerase chain reaction; ZA, zinc-adequate; ZD, zinc-deficient; ZDA, zinc-replenished.
Manuscript received December 16, 1999. Initial review completed February 8, 2000. Revision accepted May 12, 2000.
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
1.
Anant S., MacGinnitie A. J., Davidson N. O. Apobec-1, the catalytic subunit of the mammalian apo B mRNA editing enzyme, is a novel RNA-binding protein. J. Biol. Chem. 1995;270:14762-14767
2. Barnes C., Smith H. C. Apolipoprotein B mRNA editing in vitro is a zinc-dependent process. Biochem. Biophys. Res. Commun. 1993;197:1410-1414[Medline]
3. Barnes P. M., Moynahan E. J. Zinc deficiency in acrodermatitis enteropathica: multiple dietary intolerance treated with synthetic zinc. Proc. R. Soc. Med. 1973;66:327-329[Medline]
4.
Baum C. L., Teng B. B., Davidson N. O. Apolipoprotein B messenger RNA editing in the rat liver. J. Biol. Chem. 1990;265:19263-19270
5. Betts L., Xiang S., Short S. A., Wolfenden R., Carter C. W., Jr Cytidine deaminase: the 2.3 crystal structure of an enzyme: transition-state analog complex. J. Mol. Biol. 1994;235:635-656[Medline]
6.
Chen S. H., Habib G., Yang C. Y., Gu Z. W., Lee B. R., Weng S. A., Silberman S. R., Cai S. J., Deslypere J. P., Rosseneu M., Gotto A. M., Jr, Li W. H., Chan L. Apolipoprotein B-48 is the product of a messenger RNA with an organ-specific in-frame stop codon. Science (Washington, DC) 1987;238:363-366
7. Evans G. W. Zinc and its deficiency diseases. Clin. Physiol. Biochem. 1986;4:94-98[Medline]
8. Funahashi T., Giannoni F., DePaoli A. M., Skarosi S. F., Davidson N. O. Tissue-specific, developmental and nutritional regulation of the gene encoding the catalytic subunit of the rat apo B mRNA editing enzyme: functional role in the modulation of apo B mRNA editing. J. Lipid Res. 1995;36:414-428[Abstract]
9. Giannoni F., Field F. J., Davidson N. O. An improved reverse transcription-polymerase chain reaction method to study apolipoprotein gene expression in Caco-2 cells. J. Lipid Res. 1994;35:340-350[Abstract]
10. Greeve J., Altkemper I., Dieterich J.-H., Greten H., Windler E. Apolipoprotein B mRNA editing in 12 different mammalian species: hepatic expression is reflected in low concentrations of apoB-containing plasma lipoproteins. J. Lipid Res. 1993;34:1367-1383[Abstract]
11.
Hadjiagapiou C., Giannoni F., Funahashi T., Skarosi S. F., Davidson N. O. Molecular cloning of a human small intestinal apolipoprotein B mRNA editing protein. Nucleic Acids Res 1994;22:1874-1879
12.
Harris S. G., Sabio I., Mayer E., Steinberg M. F., Backus J. W., Sparks J. D., Sparks C. E., Smith H. C. Extract-specific heterogeneity in high-order complexes containing apolipoprotein B mRNA editing activity and RNA binding proteins. J. Biol. Chem. 1993;268:7382-7392
13.
Harzer G., Kauer H. Binding of zinc to casein. Am. J. Clin. Nutr. 1982;35:981-987
14. Jaccard J., Becker M. A. Statistics for the Behavioral Sciences 2nd ed. 1990:304-329 Wadsworth Publishing Belmont, CA.
15.
Koo S. I., Algilani K., Norvell J. E., Henderson D. A. Delayed plasma clearance and hepatic uptake of lymph chylomicron 14C-cholesterol in marginally zinc-deficient rats. Am. J. Clin. Nutr. 1986;43:429-437
16.
Koo S. I., Henderson D. A., Algilani K., Norvell J. E. Effect of marginal zinc deficiency on the morphological characteristics of intestinal nascent chylomicrons and the distribution of soluble apoproteins of lymph chylomicrons. Am. J. Clin. Nutr. 1985;42:671-680
17. Koo S. I., Turk D. E. Effect of zinc deficiency on intestinal transport triglyceride in the rat. J. Nutr. 1977;107:909-919
18.
Lau P. P., Chen S. H., Wang J. C., Chan L. A 40 kDa rat liver nuclear protein binds specifically to apo B mRNA around the RNA editing site. Nucleic Acids Res 1990;18:5817-5821
19.
Lau P. P., Xiong W. J., Zhu H. J., Chen S. H., Chan L. Apolipoprotein B mRNA editing is an intranuclear event that occurs posttransciptionally coincident with splicing and polyadenylation. J. Biol. Chem. 1991;266:20550-20554
20.
Lau P. P., Zhu H. J., Baldini A., Charnsangavej C., Chan L. Dimeric structure of a human apolipoprotein B mRNA editing protein and cloning and chromosomal localization of its gene. Proc. Natl. Acad. Sci. U.S.A. 1994;91:8522-8526
21. Leighton J. K., Joyner J., Zamarripa J., Deines M., Davies R. A. Fasting decreases apo B mRNA editing and the secretion of small molecular weight apo B by rat hepatocytes: evidence that the total amount of apo B secreted is regulated posttranscriptionally. J. Lipid Res. 1990;31:1663-1668[Abstract]
22. Liu G. L., Fan L. M., Redinger R. N. The association of hepatic apoprotein and lipid metabolism in hamsters and rats. Comp. Biochem. Physiol. 1991;99A:223-228
23. Nassir F., Blanchard R. K., Mazur A., Cousins R. J., Davidson N. O. Apolipoprotein B mRNA editing is preserved in the intestine and liver of zinc-deficient rats. J. Nutr. 1996;126:860-864
24. Navaratnam N., Bhattacharya S., Fujino T., Patel D., Jarmus A. L., Scott J. Evolutionary origins of apo B mRNA editing: catalysis by a cytidine deaminase that has acquired a novel RNA-binding motif as its active site. Cell 1995;81:187-195[Medline]
25.
Navaratnam N., Morrison J. R., Bhattacharya S., Patel D., Funahashi T., Giannoni F., Teng B. B., Davidson N. O., Scott J. The p27 catalytic subunit of the apo B mRNA editing enzyme is a cytidine deaminase. J. Biol. Chem. 1993a;268:20709-20712
26.
Navaratnam N., Shah R., Patel D., Fay V., Scott J. Apolipoprotein B mRNA editing is associated with UV crosslinking of proteins to the editing site. Proc. Natl. Acad. Sci. U.S.A. 1993b;90:222-226
27. Nistor A., Bulla A., Filip D. A., Radu A. The hyperlipidemic hamster as a model of experimental atherosclerosis. Atherosclerosis 1987;68:159-173[Medline]
28. O’Dell B. L., Savage J. E. Effect of phytic acid on zinc availability. Proc. Soc. Exp. Biol. Med. 1960;103:304-308
29. Powell L. M., Wallis S. C., Pease R. J., Edwards Y. H., Knott T. J., Scott J. A novel form of tissue-specific RNA processing produces apolipoprotein B-48 in intestine. Cell 1987;50:831-840[Medline]
30. Prasad A. S., Fitzgerald J. T., Hess J. W., Kaplan J., Pelen F., Dardenne M. Zinc deficiency in elderly patients. Nutrition 1993;9:218-224[Medline]
31.
Reaves S. K., Fanzo J. C., Wu J. Y. J., Wang Y. R., Wu Y., Zhu L., Lei K. Y. Plasma apolipoprotein B-48, hepatic apolipoprotein B mRNA editing and apolipoprotein B mRNA editing catalytic subunit-1 mRNA levels are altered in zinc-deficient rats. J. Nutr. 1999;129:1855-1861
32.
Reaves S. K., Hoogeveen R.C.A.J.M., Wang Y. R., Wu J.Y.J., Lei K. Y. Copper deficiency increases hepatic apolipoprotein B secretion and mRNA editing in rats. Am J. Physiol. 1996;271:C595-C604
33. Reeves P. G., Nielson F. H., Fahey G. C., Jr AIN-93 purified diets for laboratory rodents: final report of the American Institute of Nutrition ad hoc writing committee on the reformulation of the AIN-76A rodent diet. J. Nutr. 1993;123:1939-1951
34. Reinhold J. G., Faradji B., Abaidi P., Ismail-Beiji F. Decreased absorption of calcium, magnesium, zinc and phosphorus by humans due to increased fiber and phosphorus consumption as wheat bread. J. Nutr. 1976;106:493-503
35.
Roth G. J. Developing relationships: arterial platelet adhesion, glycoprotein Ib, and leucine-rich glycoproteins. Blood 1991;77:5-19
36. Sandstead H. H. Is zinc deficiency a public health problem?. Nutrition 1995;11:87-92[Medline]
37. Scanu A. M., Fless G. M. Lipoprotein (a): heterogeneity and biological relevance. J. Clin. Investig. 1990;85:1709-1715
38.
Shah R. R., Knott T. J., Legros J. E., Navaratnam N., Greeve J. C., Scott J. Sequence requirements for the editing of apo B mRNA. J. Biol. Chem. 1991;266:16301-16304
39.
Smith H. C., Kuo S. R., Backus J. W., Harris S. G., Sparks C. E., Sparks J. D. In vitro apolipoprotein B mRNA editing: identification of a 27S editing complex. Proc. Natl. Acad. Sci. U.S.A. 1991;88:1489-1493
40. Spady D. K., Dietschy J. M. Interaction of dietary cholesterol and triglycerides in the regulation of hepatic low density lipoprotein transport in the hamster. J. Clin. Investig. 1988;81:300-309
41.
Teng B. B., Burant C. F., Davidson N. O. Molecular cloning of an apolipoprotein B messenger RNA editing protein. Science (Washington, DC) 1993;260:1816-1818
42.
Teng B. B., Davidson N. O. Evolution of intestinal apolipoprotein B mRNA editing. J. Biol. Chem. 1992;267:21265-21272
43.
Teng B. B., Verp M., Salomon J., Davidson N. O. Apolipoprotein B messenger RNA editing is developmentally regulated and widely expressed in human tissues. J. Biol. Chem. 1990;265:20616-20620
44.
Wu J.Y.J., Reaves S. K., Wang Y. R., Wu Y., Lei P. P., Lei K. Y. Zinc deficiency decreases plasma level and hepatic mRNA abundance of apolipoprotein A-I in rats and hamsters. Am. J. Physiol. 1998;275:C1516-C1525
45.
Yamanaka S., Poksay K. S., Balestra M. E., Zeng G.-Q., Innerarity T. L. Cloning and mutagenesis of the rabbit apoB mRNA editing protein. J. Biol. Chem. 1994;269:21725-21734
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