Osmotic Regulation Section, Laboratory of Kidney and Electrolyte Metabolism, National Heart, Lung and Blood Institute, National Institutes of Health, Bethesda, Maryland 20892-1603
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ABSTRACT |
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The molecular
mechanisms underlying adaptation to hyperosmotic stress through the
accumulation of organic osmolytes are largely unknown. Yet, among
organisms, this is an almost universal phenomenon. In mammals, the
cells of the renal medulla are uniquely exposed to high and variable
salt concentrations; in response, renal cells accumulate the osmolyte
sorbitol through increased transcription of the aldose reductase (AR)
gene. In cloning the rabbit AR gene, we found the first evidence of an
osmotic response region in a eukaryotic gene. More recently, we
functionally defined a minimal essential osmotic response element (ORE)
having the sequence CGGAAAATCAC(C) (bp 1105 to
1094). In
the present study, we systematically replaced each base with every
other possible nucleotide and tested the resulting
sequences individually in reporter gene constructs. Additionally, we categorized hyperosmotic response by electrophoretic mobility shift assays of a 17-bp sequence (
1108 to
1092)
containing the native ORE as a probe against which the
test constructs would compete for binding. In this manner, binding
activity was assessed for the full range of osmotic responses obtained.
Thus we have arrived at a functional consensus for the mammalian ORE,
NGGAAAWDHMC(N). This finding should accelerate the discovery of genes
previously unrecognized as being osmotically regulated.
osmoregulation; osmotic stress; organic osmolytes; gene regulation; gene expression
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INTRODUCTION |
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EVOLUTIONARILY, ADAPTATION to hyperosmotic stress through the accumulation of osmotically active organic solutes (organic osmolytes) is a highly conserved mechanism. In organisms other than halophilic bacteria, cells accumulate high concentrations of organic osmolytes instead of inorganic ions. In cells that respond with an initial short-term adaptive accumulation of inorganic ions, inorganic ions are replaced by organic osmolytes in the long term. This occurs because elevated concentrations of organic osmolytes apparently do not perturb the macromolecular function as would equivalent concentrations of inorganic ions (28). Although the cellular responses to hyperosmotic stress are among the most profound, the molecular mechanisms involved have only begun to be addressed. Thus, in contrast to what is known about another environmental perturbation, heat shock, relatively little is known about signal transduction between the initial extracellular stimulus (hyperosmolality) and the ultimate adaptive response (13, 23, 27).
Hyperosmotic accumulation of organic osmolytes is transcriptionally regulated. Organic osmolytes include betaine in bacteria (e.g., Escherichia coli) (17), glycerol in yeast (e.g., Saccharomyces cerevisiae) (1, 14), and sorbitol and betaine in cells of the mammalian renal medulla, which, among mammalian cells, are uniquely exposed to hyperosmotic stress in normal physiological conditions (9). In bacteria, transport of betaine is encoded by ProU (17). Osmotic control of proU operon transcription is exerted for the most part by a negative-acting "downstream regulatory element" (reviewed in Refs. 11, 17, and 20). There additionally may be minimal involvement of an "upstream activating region" extending ~200 bp in length (17). Although deletion analysis of this region has been performed, no discrete cis element that confers osmotic response has been identified (16). In S. cerevisiae, GPD1 encodes glycerol-3-phosphate dehydrogenase, which catalyzes the synthesis of glycerol. Osmotic stress increases glycerol-3-phosphate dehydrogenase activity and mRNA levels (1), but osmotically responsive cis elements in the GPD1 gene have not been identified. In the yeast high-osmolarity glycerol (HOG) response, most is known about the cascade of signals that immediately follows an increase in osmolarity (reviewed in Ref. 14). HOG1 (a mitogen-activated protein kinase gene)-dependent osmotic induction of genes other than GPD1 has been shown to act via yeast stress response elements (STREs) (22). Yeast STREs function under various insult conditions, not only osmotic stress, and there is no homology between the osmotic response element (ORE) and the STREs.
In mammalian renal medullary cells, including cultured PAP-HT25 cells, which are derived from the rabbit inner medulla, hyperosmotic stress results in the accumulation of sorbitol as a predominant osmolyte (2, 3). Sorbitol accumulates because of a rise in the rate of synthesis of aldose reductase (AR) (19), the enzyme that catalyzes the conversion of glucose to sorbitol. Using PAP-HT25 cells, we originally demonstrated that hyperosmotic stress increases the transcription of the AR gene (24), which leads to a rise in AR mRNA levels (10). In cloning and characterizing the rabbit AR gene, we found the first evidence of a eukaryotic ORE (8).
Since then, there have been several discoveries of mammalian OREs,
including the definition of the minimal essential ORE of the rabbit AR
gene (7). A sequence containing an ORE was identified for the canine
Na+- and
Cl-coupled betaine
transporter gene (25), within which it is possible to find homology
with the rabbit AR gene minimal essential ORE. Another
functional ORE, which is now designated ORE-X, was originally thought
to be upstream of the human AR gene (21). ORE-X is strongly homologous
(9 of 11 bp are identical) to the rabbit AR gene ORE, but the gene to
which it belongs is as yet unknown (12). Putative OREs of the AR gene
in the human (12) and mouse (4) now have been found. This constitutes a
total count of five OREs, three of which are in the same gene (the AR
gene), although the genes are from different species. The scarcity of
demonstrated active OREs makes it unlikely that an accurate, strong
consensus for mammalian minimal essential OREs can be derived merely by
sequence comparison. Here, we present the systematic
derivation by functional assessment of the mammalian minimal essential ORE.
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METHODS |
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Reporter gene expression analysis of transient transfectants.
Expression vector ARL has been described previously (7). ARL contains
the rabbit AR gene promoter (bp 208 to +27) in unique Xho
I-Hind III sites immediately upstream
of the firefly luciferase gene. ARL was previously
demonstrated to exhibit basal promoter activity (7). pRLB19 was
constructed by subcloning the B19 promoter (8), a strong,
constitutively active promoter from the B19 parvovirus, into the
Xho
I-Hind III sites of the multiple cloning region immediately upstream of the
Renilla luciferase gene in the
promoterless pRL-null vector (Promega).
Transfection and luciferase assays. Transfections were performed as previously described (7, 8). Briefly, rabbit renal medulla PAP-HT25 cells (passage 67-83) were grown in isosmotic medium (300 mosmol/kgH2O) (26) in 150-mm-diameter dishes (Corning) and cotransfected with a given ARL firefly luciferase construct (3 µg) and pRLB19 (0.03 µg) by using a CellPhect transfection kit (Pharmacia). From each transfected 150-mm-diameter dish, cells were seeded into six 35-mm-diameter dishes and left overnight. The medium in three of the dishes was changed to fresh isosmotic medium (300 mosmol/kgH2O); the medium in the other three dishes was changed to the same medium made hyperosmotic (500 mosmol/kgH2O) with NaCl. Twenty-four hours after changing the medium, cells were harvested by adding 100 µl of passive lysis buffer (Dual-Luciferase reporter assay system; Promega).
Cell lysates were analyzed for total protein and firefly and Renilla luciferase activities [in relative light units (RLU)] by using the following kits in accordance with the manufacturers' instructions. Total protein was determined by using the Bio-Rad protein assay kit withTransfection data analyses. Firefly luciferase activity (from the experimental ARL construct) was normalized by Renilla luciferase activity (from the cotransfected pRLB19 construct; both activities were in RLU/µg of cell protein). Osmotic response was calculated as the hyperosmotic-to-isosmotic ratio, i.e., the ratio of the firefly luciferase to the Renilla luciferase activity measurements under the two conditions. Osmotic response ratios were compared by a one-way ANOVA on square root-transformed ratio data followed by Dunnett's multiple-comparison test for separation of significant means.
Electrophoretic mobility shift assays.
Nuclear protein extracts were prepared from PAP-HT25 cells maintained
in isosmotic medium (300 mosmol/kgH2O) and from cells exposed for 18-24 h to the same medium made hyperosmotic (500 mosmol/kgH2O) with NaCl (10).
Nuclear protein extracts were prepared by the method of Dignam et al.
(5, 6) with buffer C containing 0.42 M KCl.
32P-end-labeled double-stranded
oligonucleotides (bp 1108 to
1092 in the rabbit AR gene
containing the minimal essential ORE,
1105 to
1094; 100 fmol) were incubated with 6 µg nuclear protein extract, 0.5 µg
poly(dI-dC), and 0-5 pmol (0- to 50-fold molar excess) unlabeled
specific (AR gene bp
1108 to
1092) or mutant (AR gene bp
1108 to
1092 or
1105 to
1094 containing
single-base substitutions) competitor in 10 µl of binding buffer (21)
for 30 min at room temperature. All oligonucleotides were synthesized
directly without restriction enzyme sites at their 5' or 3'
ends. Reaction products were separated by electrophoresis on a 4%
polyacrylamide gel (30:0.8, acrylamide-bisacrylamide) in 0.5×
Tris-borate-EDTA buffer at 4°C (21). Autoradiograms were exposed at
80°C. The electrophoretic mobility shift assay (EMSA) was
repeated at least twice for each construct examined.
Densitometric analysis. A densitometric analysis was performed on the shifted bands, avoiding the nonspecific signal that was sometimes present at the lane margins. The densitometric value of the band(s) shifted in the presence of hyperosmotic extract was used as the maximum binding capacity. This value was typically three to four times that of the equivalent band(s) shifted in the presence of isosmotic extract. The binding capacity after a single-base substitution was determined by the ability of an excess (10 or 50 times the mass of the probe) of these mutants to compete effectively, that is, to significantly reduce the maximum binding capacity of the probe in the presence of hyperosmotic extract, as described by Ruepp et al. (21). Significant reduction was equated with the ability of a 50-fold excess of a mutant to reduce binding capacity at least to the level found in the presence of isosmotic extract.
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RESULTS AND DISCUSSION |
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The original demonstration of a putative ORE was in a 3,221-bp fragment
of the 5' flanking region (3429 to
209) of the
rabbit AR gene (8). Subsequently, we functionally defined, within that
fragment, the smallest sequence capable of conferring osmotic response
on a downstream gene, a minimal essential ORE (7). The ability of this
ORE to confer osmotic response was independent of that of other
putative cis elements that may
potentiate the response (7).
We derived the putative consensus for a mammalian ORE (Fig.
1) from the minimal essential
rabbit AR gene ORE (7), putative AR gene OREs from humans (12) and mice
(4), ORE-X from an unknown gene (12, 21), and the ORE of the canine
Na+- and
Cl-coupled betaine
transporter gene (TonE) (25). Three of these are in the same gene, and
a set of only three different genes is insufficient to clearly define a
true consensus. Therefore, we have now functionally defined a consensus
for mammalian OREs. Starting with the rabbit AR gene minimal essential
ORE (
1105 to
1094) (Fig. 1), we systematically replaced
each base individually with every other possible nucleotide and tested
each construct. This allowed us to examine the nucleotide requirement
at each position within the minimal essential ORE for the ability to
confer osmotic response to a firefly luciferase gene driven by the AR promoter (
208 to +27) (7, 8) in transient transfection assays
(Fig. 1). Relative to the osmotic response of the negative control (ARL
containing the promoter alone), we defined the ability to confer an
osmotic response as a hyperosmotic-to-isomotic ratio of >1.4 based on
statistical analysis (square root transformation of ratio data followed
by one-way ANOVA with separation of significant means by Dunnett's
multiple-comparison test). Additionally, to assess binding activity, we
used EMSAs. In these, a 17-bp sequence (
1108 to
1092)
containing the native ORE (
1105 to
1094) was used as a
probe against which the test constructs would compete for binding (Fig.
2). We initially determined the binding
profiles of the majority (8 of 9) of constructs that were clearly
positive in osmotic response (ratio
1.7) and those that were clearly negative (14 of 24) in osmotic response (ratio
1.3). The binding activity of constructs with ratios
1.7 was positive; the binding of
constructs with ratios
1.3 was negative. All other constructs having
intermediate response ratios (1.4-1.6) were then examined. In this
manner, binding activity was assessed for the full range of osmotic
response ratios obtained (2.7-0.8; Fig. 1). EMSA was repeated at
least twice for each construct examined. Constructs that did not
effectively compete (i.e., binding capacity reduced to the level found
in the presence of isosmotic extract or lower), even at a 50-fold molar
excess, with the native probe were considered osmotically inactive
(Fig. 2). This also defined construct hyperosmotic-to-isosmotic ratios
of >1.4 as active (Fig. 1). Accordingly, the nucleotide requirements
for each position within the minimal essential ORE were defined as
follows. Position
1105 can be filled by any of the four
nucleotides. However, position
1105 cannot be eliminated and the
osmotic response retained (7). Positions
1104 through
1100 must be GGAAA, respectively. Positions
1099 through
1096 can vary;
1099 can be A or T;
1098 can be T,
A, or G;
1097 can be C, A, or T; and
1096 can be A or C. Position
1095 must be C, and
1094 can be any of the four
nucleotides. These results define the functional consensus for the
mammalian ORE as NGGAAA(A/T)(T/A/G)(C/A/T)(A/C)C(N) or,
in single-letter nomenclature, as NGGAAAWDHMC(N). The
further dispensability of G nucleotides in each strand has been
addressed by Miyakawa et al. (18) by using an in vivo footprinting
technique in Madin-Darby canine kidney (MDCK) cells exposed to
hypertonic medium. They concluded that G residues in the canine betaine
transporter gene ORE corresponding to positions
1104,
1103, and
1095 (in the rabbit AR gene ORE) are
indispensable, whereas a G at position
1098 is not. These
results are consistent with the functional consensus ORE. However,
because this functional consensus was derived from the known mammalian
OREs, which have strong homology, the possibility of the future
discovery of mammalian OREs that bear no resemblance to this consensus
cannot be discounted.
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Miyakawa et al. (18) recently reported deriving a consensus ORE (Fig.
3). Their consensus differs from ours. A
principal difference is due to the fact that Miyakawa et al. did not
derive their consensus entirely empirically. For example, none of the bases were individually replaced by all other possible nucleotides. Therefore, many of the conclusions are derived from assumptions; e.g.,
pyrimidines (C or T) and purines (A or G) were considered equal without
consideration of differences in hydrogen bonding (A and T form two
bonds; C and G form three bonds). Specific examples follow (Fig. 3). 1) The base
corresponding to position 1099 of the rabbit AR gene ORE was
concluded to be N, whereas we conclude that it can be an A or T. It is
an A in all known active OREs, and Miyakawa et al. replaced it only
with a T and found a positive response. They concluded that this base
could be an N simply because a pyrimidine (T) and a purine (A) had been
tested. This conclusion did not take into account the fact that both A
and T form two hydrogen bonds. We demonstrated
empirically that neither C nor G can effectively substitute at this
position. 2) The base corresponding to position
1095 of the rabbit AR gene ORE was concluded to be a
C or T, whereas we conclude that it can only be a C. Miyakawa et al.
based their conclusion on the presence of a T in OreA and OreB from the
human AR gene "osmotic response region" (12) and did not test it
empirically. Neither OreA nor OreB has been shown to be
independently active. Independent osmotic activity is the underlying
assumption in the experimental approach of Miyakawa et al. (and of
ours). However, two notable discrepancies in osmotic response were
found when individual bases were substituted at AR gene ORE positions
1105 and
1100. We found osmotic response with a G at
1105, whereas Miyakawa et al. did not. Miyakawa et al. found
osmotic response with a T at
1100 and concluded that
1100
could be an N. We substituted at
1100 with C, G, and T and lost
osmotic response, concluding that the base at
1100 could only be
an A. The reasons for these discrepancies are not apparent; there is,
however, at least one important experimental difference. Miyakawa et
al. used a tandem repeat of the OREs being tested, whereas we used a
single copy. We do not believe these discrepancies to be a result of a
species difference (dog vs. rabbit cell lines) because we obtained
osmotic response with the AR gene ORE in MDCK cells (as Miyakawa et al.
used) and with TonE in PAP-HT25 cells (unpublished results).
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On the basis of our data, we can conclude that native, active OREs fit
this functional consensus. However, we then asked whether the converse
was true, that is, is any sequence that fits the functional consensus
ORE necessarily osmotically responsive? The functional consensus allows
more latitude per position than the predicted putative consensus
derived merely by maximizing the homology of presently known native
OREs (Figs. 1 and 4). We asked how many
substitutions that fit the functional consensus but deviate from
maximum homology among known native OREs can one make and still obtain
osmotic response. Figure 4 illustrates one example of this test. When
the test construct contained either four or three positions that
deviated from maximum homology, osmotic response was lost, although the
position changes still fit the functional consensus (Fig. 4). Two or
fewer position changes that deviated from maximum homology resulted in
the retention of osmotic response. These data are consistent with
previous observations that most functional elements must be related to
the consensus by no more than one or two substitutions (15). We
conclude that, although a functional ORE fits the functional consensus,
the converse is not true. Any gene sequence identified as a possible
ORE by homology scanning with the consensus sequence must ultimately be
assessed by the expression of osmotic response. For example, a putative osmotic response region in the bovine
Na+/myo-inositol
cotransporter gene (SMIT) was recently identified (29). This region
contains a sequence that shares partial similarity with that of the ORE
(7 of 11 bp are identical); however, the sequence has G at
corresponding base 1099, and this does not fit the functional
consensus. Whether the 11-bp SMIT ORE is osmotically responsive remains
to be examined. Our determination of a functional consensus for
mammalian OREs should accelerate the recognition of genes that are
osmotically regulated.
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. §1734 solely to indicate this fact.
Address for reprint requests: J. D. Ferraris, National Institutes of Health, Bldg. 10, Rm. 6N260, 10 Center Dr., MSC 1603, Bethesda, MD 20892-1603 (E-mail: jdf{at}helix.nih.gov).
Received 2 September 1998; accepted in final form 24 November 1998.
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