1 Department of Pharmacology and Toxicology and 2 Department of Pharmaceutical Sciences, University of Arkansas for Medical Sciences, Little Rock, Arkansas 72205-7199
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Collagen IV is found in the renal proximal tubular cell (RPTC) basement membrane and is a mediator of renal development and function. Pharmacological concentrations of L-ascorbic acid phosphate (AscP) promote the repair of physiological functions in RPTC sublethally injured by S-(1,2-dichlorovinyl)-L-cysteine (DCVC). We hypothesized that AscP promotes RPTC repair by stimulating collagen IV synthesis and/or deposition. RPTC exhibited increased synthesis but decreased deposition of collagen IV after DCVC exposure. In contrast, RPTC cultured in pharmacological concentrations of AscP maintained collagen IV deposition. The activity of prolyl hydroxylase was decreased in RPTC after DCVC injury, an effect that was partially attenuated in injured RPTC cultured in pharmacological concentrations of AscP. The addition of exogenous collagen IV to the culture media of DCVC-injured RPTC promoted the repair of mitochondrial function and Na+-K+-ATPase activity. However, neither collagen I, laminin, nor fibronectin promoted cell repair. These data demonstrate an association between AscP-stimulated deposition of collagen IV and exogenous collagen IV and repair of physiological functions, suggesting that collagen IV plays a specific role in RPTC repair after sublethal injury.
cell injury; regeneration; extracellular matrix; collagen synthesis and deposition; prolyl hydroxylase
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
THE MESHLIKE BASEMENT
MEMBRANE (BM) provides structural support and influences the
growth, function, and survival of many cell types in most organ systems
(15). Collagens are extracellular matrix (ECM) proteins
that form the renal tubular BM with other ECM proteins such as laminin
and heparan sulfate proteoglycans (29). The most abundant
type of collagen in the BM of the glomerulus and renal tubules is
collagen IV, a globular, nonfibrillar protein. This characteristic
distinguishes it from collagen I, the major fibrillar component of
connective tissues and the second most abundant ECM protein in the
proximal tubular BM (12, 21). Collagen IV forms a
triple-helical monomer that consists most often of two 1(IV) chains
and one
2(IV) chain or three
1(IV) chains (14, 42).
The collagen IV chains
3(IV),
4(IV),
5(IV), and
6(IV) have
been identified and can associate in various combinations (15,
20). However, these isoforms have not been detected in the human
proximal tubule or in primary cultures of rabbit renal proximal tubular
cells (12, 29). Except in rodents, their renal expression
appears to be limited to the distal tubular BM and the glomerular BM,
where they have been implicated in the development of Goodpasture and
Alport syndromes and diffuse leiomyomatosis (11, 15, 19, 20, 27,
29). By using functional analyses of cell-matrix interactions,
collagen IV has been shown to play a crucial role in tubular function
and kidney development (31). Because collagen IV is an
important anchorage substrate for many cell types, especially in the
kidney, the regulation of collagen IV synthesis and degradation plays
an important role in cell function, growth, migration, and organ
remodeling (15).
Under conditions of ischemia or after acute chemical exposures, renal epithelial cells may die or detach from the ECM and slough into the tubular lumen. Here, they may aggregate with other sloughed cells, forming casts that cause tubular obstruction. Cells that do not die or that become detached from the ECM are thought to dedifferentiate, proliferate, and migrate to denuded areas of the tubule, thus replacing the sloughed cells. The cells of the newly lined tubule may then differentiate, promoting the return of normal tubular function and overall renal function (1). The roles of collagens and other ECM proteins in renal cell survival, migration, and function have been examined (4). Surprisingly, few reports exist regarding the role of collagens in cellular repair and regeneration, although proliferation, migration, and return of normal function do contribute to renal regeneration after injury (49).
Ascorbic acid is known to prevent the effects of scurvy, a disease
characterized by defective connective tissue resulting from decreased
collagen synthesis (40). In posttranslational processing
mechanisms, ascorbate acts as an essential iron-reducing cofactor in
the production of collagens, specifically in the hydroxylation of
susceptible proline and lysine residues in procollagen chains. These hydroxylation reactions are catalyzed by prolyl and lysyl hydroxylases, respectively, and are necessary for the proper folding of
procollagen triple helices as well as other posttranslational modifications, including glycosylation and monomer cross-linking (9, 24). Insufficiently hydroxylated procollagens have
been shown to accumulate intracellularly, be deposited much more
slowly, and be targeted for rapid degradation both intracellularly and extracellularly (17, 18, 42). Ascorbic acid also is known to promote the synthesis of both fibrillar and nonfibrillar collagen types in an array of cell types in vitro (10, 13, 33, 45). In addition, ascorbate has been suggested to act pretranslationally by
stimulating mRNA expression of multiple collagen types in various culture systems, independent of its role as an enzymatic cofactor (6, 14, 32, 41, 46). Ascorbic acid has been implicated as
an important mediator of cell growth and differentiation in a variety
of cell types, through its effects on collagen synthesis and deposition
(2). Through mechanisms unrelated to ECM production, ascorbic acid has been shown to both stimulate and inhibit cell proliferation depending on ascorbate concentration and cell type (7, 16, 48). Previous studies from our laboratory
demonstrated that ascorbic acid promotes increased cell growth and
density and improvement of key physiological functions, including
brush-border enzyme activity, basal oxygen consumption
(QO2), and
Na+-K+-ATPase activity in primary cultures of
rabbit renal proximal tubular cells (RPTC) (39).
The halocarbon conjugate S-(1,2-dichlorovinyl)-L-cysteine (DCVC) is a model toxicant that produces RPTC necrosis and acute renal failure (23). We previously showed that primary cultures of rabbit RPTC sublethally injured by DCVC neither proliferate nor repair physiological functions (37). In those experiments, RPTC were grown under physiological concentrations of all culture media supplements including 50 µM L-ascorbic acid 2-phosphate (AscP). However, on addition of pharmacological concentrations of AscP (500 µM), RPTC exposed to DCVC were able to proliferate and repair physiological functions, although AscP provided no protective effect during injury. In addition, pharmacological concentrations of ascorbic acid were shown to stimulate collagen IV synthesis and deposition in uninjured RPTC (36). Therefore, the goals of this study were 1) to examine the effect of DCVC on the synthesis, deposition, and proline hydroxylation of collagen IV in DCVC-injured RPTC, 2) to determine whether AscP-stimulated synthesis and/or deposition of collagen IV is associated with AscP-stimulated repair of physiological functions in DCVC-injured RPTC, and 3) to characterize the specific role of collagen IV and other ECM proteins in RPTC repair.
![]() |
MATERIALS AND METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Materials. Female New Zealand White rabbits (1.5-2.0 kg) were purchased from Myrtle's Rabbitry (Thompson Station, TN). DCVC was a generous gift from Dr. T. W. Petry (Pharmacia Upjohn, Kalamazoo, MI) and was synthesized as previously described (30). AscP (magnesium salt) was purchased from Wako Chemicals (Richmond, VA). Ouabain was obtained from RBI/Sigma (Natick, MA). L-[35S]methionine (>1,000 Ci/mmol) was purchased from ICN Biomedicals (Costa Mesa, CA). L-[14C]proline (0.275 Ci/mmol) and L-[4-3H]proline (24 Ci/mmol) were purchased from New England Nuclear Life Science Products (Boston, MA). The mouse anti-collagen IV monoclonal antibody M3F7, developed by Dr. Heinz Furthmayr, was obtained from the Developmental Studies Hybridoma Bank (University of Iowa, Iowa City, IA). Hyperfilm enhanced chemiluminescence was purchased from Amersham Life Science (Cleveland, OH). Hyamine hydroxide was purchased from ICN Radiochemicals (Irvine, CA). Rat tail collagen I was purchased from Collaborative Biochemical Products (Bedford, MA). Human cellular fibronectin was purchased from Upstate Biotech (Lake Placid, NY). Human placenta collagen IV, laminin, and all other materials were purchased from Sigma (St. Louis, MO).
Isolation of proximal tubules and culture conditions.
Rabbit renal proximal tubules were isolated by using the iron oxide
perfusion method and grown in 35-mm tissue culture dishes or 48-well
cell culture clusters under improved conditions as previously described
(38, 39). The cell culture medium was a 1:1 mixture of
DMEM/Ham's F-12 (without D-glucose, phenol red, or sodium
pyruvate) supplemented with 15 mM HEPES buffer, 2.5 mM
L-glutamine, 1 µM pyridoxine HCl, 15 mM sodium
bicarbonate, and 6 mM lactate. Hydrocortisone (50 nM), selenium (5 ng/ml), human transferrin (5 µg/ml), bovine insulin (10 nM), and AscP (50 or 500 µM) were added to fresh culture medium immediately before
daily change of media. AscP was used because L-ascorbic acid is unstable in culture media. AscP is stable in culture media for
7 days at 37°C and, after intracellular dephosphorylation, has the
same effect on cultured cells as L-ascorbic acid
(13).
Sublethal injury of RPTC.
Confluent monolayers of RPTC (day 6 after seeding) were
exposed to 200 µM DCVC (dissolved in water) for 1.75 h followed
by toxicant removal and addition of fresh culture media. This method produces ~50% cell death and loss 24 h after exposure. On
days 1, 4, and 6 after DCVC exposure,
the ability of the remaining RPTC to regenerate and repair
physiological function was monitored as described below. In some
experiments, immediately after DCVC exposure and after daily change of
media through day 6 after injury, collagen I (0, 5, 15, or
50 µg/ml), collagen IV (0, 5, 15, or 50 µg/ml), laminin (50 µg/ml), or cellular fibronectin (50 µg/ml) was added to the culture
media of uninjured and DCVC-injured RPTC grown in physiological
concentrations of AscP. Exogenous collagens I and IV used in this study
were triple helical and contained only 1 and
2 chains in
conformations similar to those found in renal BM (3, 25).
Laminin was a mixture of biologically active laminin chains found in
most epithelial tissues (8, 50), and cellular fibronectin
was composed of functional dimers (51).
Cell density. Measurement of monolayer protein content over time was used to estimate cell density. On days 1, 4, and 6 after sublethal DCVC injury, RPTC monolayers were washed with PBS and solubilized in Triton buffer (0.05% Triton X-100, 100 mM Tris-base, 150 mM NaCl, pH 7.5). Samples were sonicated, and protein concentrations were determined by using the method of Lowry (26) or the bicinchoninic acid microassay according to the manufacturer's instructions (Pierce, Rockford, IL).
Basal oxygen consumption. On days 1, 4, and 6 after sublethal DCVC injury, RPTC bathed in 37°C culture medium were gently detached from culture dishes with a rubber policeman and transferred to a 37°C QO2 chamber. Basal RPTC QO2 was measured polarographically by using a Clark-type electrode as described previously (39).
Na+-K+-ATPase
activity.
On days 1 and 6 after sublethal DCVC injury,
total ATPase activity was measured using a modification of a previously
described procedure (44). Briefly, RPTC cultured in
48-well cell culture clusters were scraped and incubated in
dissociation buffer [5 mM HEPES (pH 7.4), 25 mM imidazole, 1% BSA,
0.065% SDS] for 10 min at room temperature and placed on ice. The
dissociated RPTC were then diluted fivefold with an additional
dissociation buffer minus SDS. Aliquots of dissociated RPTC were
combined with fresh ATPase assay buffer [(in mM) 2.54 MgCl2, 100 NaCl, 10 KCl, 5 HEPES, 2.54 Na2ATP,
2.54 phospho(enol)pyruvate, and 0.5 -NADH], as well as 10 U/ml lactate dehydrogenase and 7 U/ml pyruvate kinase. ATPase activity was measured spectrophotometrically under linear conditions as
the oxidation of
-NADH to NAD+ at 37°C in the absence
or presence of ouabain (0.1 mM) at a wavelength of 340 nm.
Na+-K+-ATPase activity was calculated as total
ATPase activity minus ouabain-insensitive ATPase activity.
Immunoprecipitation of synthesized and deposited collagen IV.
Immunoprecipitation of newly synthesized and deposited collagen IV was
performed by using the method of Niki et al. (34), with
some modifications. On days 1, 4, and
6 after sublethal DCVC injury, RPTC were metabolically
labeled for 24 h with 25 µCi/ml of
L-[35S]methionine. Cell monolayers were washed with
PBS, solubilized for 30 min in lysis buffer [20 mM Tris · HCl
(pH 8.8), 2 mM EDTA, 0.2 mM phenylmethylsulfonyl fluoride (PMSF), 10 mM
N-ethylmaleimide, 1% Triton X-100, 1% sodium deoxycholic
acid, 150 mM NaCl, and 1 mM EGTA] containing protease inhibitors (25 µg/ml pepstatin A, 12.5 µg/ml leupeptin, 12.5 µg/ml aprotinin),
and centrifuged at 15,000 g for 10 min at 4°C. The
resulting supernatants (soluble fraction, cell-associated collagen)
were transferred to fresh tubes, snap-frozen in liquid N2,
and stored at 80°C. Pelleted material (insoluble fraction,
ECM-associated collagen) was resuspended in pellet solubilization
buffer [(in mM) 20 Tris · HCl (pH 8.8), 2 EDTA, 0.2 PMSF, 10 N-ethylmaleimide, and 10 dithiothreitol, as well as 1%
SDS], sonicated, boiled for 5 min, and incubated in the presence of
iodoacetamide (25 mM) for 30 min at 37°C with shaking. Samples were
centrifuged at 15,000 g for 10 min at 4°C. Supernatants
were then transferred to fresh microcentrifuge tubes, snap-frozen in
liquid N2, and stored at
80°C.
SDS-PAGE and autoradiography.
Radiolabeled collagen IV was separated by SDS-PAGE and visualized by
autoradiography (22). Each radiolabeled collagen IV immunoprecipitate (20 µl) was subjected to electrophoresis, and the
gels were stained, dried, and exposed to film for 3 wk at 80°C.
After film development, densitometry of the resulting 206-kDa bands was
calculated using National Institutes of Health Image software.
Proline hydroxylation in newly synthesized collagen IV. Proline hydroxylation in collagen molecules occurs at the 4-trans position. Therefore, the extent of proline hydroxylation can be estimated by the loss of hydrogen from the 4-trans position of proline in the hydroxylation reaction. We modified a previously described dual-labeling technique for the determination of proline hydroxylation by using L-[14C]proline and L-[4-3H]proline incorporation into collagen IV and measuring the ratio of 3H to 14C in collagen IV molecules (5). A greater extent of proline hydroxylation at the 4-trans position of proline will result in a decrease in the ratio of 3H to 14C in newly synthesized collagen molecules due to the loss of 3H. RPTC were dual labeled for 24 h with L-[14C]proline (15 µCi/ml) and L-[4-3H]proline (62.5 µCi/ml) immediately after DCVC exposure. Cell-associated and deposited proteins were harvested, and collagen IV was immunoprecipitated as described above. Immunoprecipitated collagen IV was visualized by SDS-PAGE as mentioned above, the 206-kDa bands in the destained gels were excised, and the protein was extracted with 90% hyamine hydroxide overnight at 37°C. [14C]proline and [4-3H]proline in collagen IV was determined by using liquid scintillation spectrometry, and the extent of proline hydroxylation was calculated as the ratio of 3H to 14C.
Statistical analysis. RPTC isolated from one rabbit represent one experiment (n = 1) that consists of data collected from two to six plates of cells. Experiments were repeated until an n of three to five was reached. Data are presented as means ± SE. Significant differences between treatment groups (P < 0.05) were determined by using SigmaStat one-way ANOVA or two-way ANOVA as necessary and Student-Newman-Keuls tests for the comparison of multiple means (Jandel Scientific, San Rafael, CA).
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Cell density.
Monolayer protein content was used to measure cell density over a 6-day
recovery period in RPTC sublethally injured by DCVC. There were no
significant differences at any time point in monolayer protein content
between untreated RPTC grown in 50 or 500 µM AscP. One day after DCVC
exposure (200 µM), RPTC cultured in 50 µM AscP exhibited a 56%
decrease in monolayer protein content compared with control,
representing cell death and loss (Fig.
1). RPTC cultured in the presence of 500 µM AscP sustained the same degree of cell death and loss (55%) on
day 1. Monolayer protein content 4 days after DCVC exposure
remained at day 1 levels in RPTC incubated in 50 or 500 µM
AscP. However, 6 days after DCVC exposure, monolayer protein content in
DCVC-treated RPTC that was cultured in 500 µM AscP was 34% greater
than that of DCVC-treated RPTC grown in 50 µM AscP. These data show
that pharmacological concentrations of ascorbic acid do not protect
against DCVC-induced cell injury and loss but do promote an increase in
cell density over time in RPTC sublethally injured by DCVC.
|
Basal QO2 and
Na+-K+-ATPase
activity.
Basal QO2 (a measure of mitochondrial function)
and Na+-K+-ATPase activity were used as
measures of physiological functions in RPTC over the 6-day recovery
period after DCVC exposure. Basal QO2 and
Na+-K+-ATPase activity in untreated RPTC grown
in 50 and 500 µM AscP were equivalent at all time points. One day
after DCVC exposure (200 µM), RPTC grown in both 50 and 500 µM AscP
exhibited decreases in QO2 and
Na+-K+-ATPase activity of ~50 and 65%,
respectively (Fig. 2, A and
B). QO2 remained at day 1 levels on day 4 after DCVC exposure in RPTC incubated in 50 or 500 µM AscP. On day 6, QO2 in
RPTC grown in 500 µM AscP was 49% higher than that in RPTC grown in
50 µM AscP and equal to that in controls. Similarly,
Na+-K+-ATPase activity on day 6 was
50% higher in DCVC-injured RPTC cultured in 500 µM AscP than in
injured RPTC cultured in physiological concentrations of AscP. These
data show that pharmacological concentrations of ascorbic acid do not
prevent the inhibition of physiological functions caused by DCVC
exposure but do stimulate physiological repair in RPTC sublethally
injured by DCVC.
|
Synthesis of collagen IV.
Figure 3 shows a representative
autoradiograph of newly synthesized but not deposited
35S-labeled collagen IV in RPTC after exposure to DCVC.
Densitometric analysis of the 206-kDa collagen IV band was performed on
scanned images of individual autoradiographs from four separate
experiments (Fig. 4). To illustrate
changes in collagen IV synthesis over time in each group, values are
expressed as a percentage of day 1 controls grown in 50 µM
AscP. Collagen IV synthesis on day 1 was decreased in
uninjured RPTC grown in 500 µM AscP compared with those grown in 50 µM AscP. One day after DCVC exposure, there was an equivalent
1.8-fold increase in collagen IV synthesis in RPTC grown in both 50 and
500 µM AscP. Levels of collagen IV synthesis at days 4 and
6 after DCVC exposure and in controls were reduced compared
with day 1 in RPTC grown in both 50 and 500 µM AscP. No
significant differences between groups were found on days 4 and 6. These findings demonstrate that RPTC sublethally
injured by DCVC increase collagen IV synthesis 1 day after the injury. However, this stimulation is independent of the concentration of
ascorbic acid.
|
|
Deposition of collagen IV.
Figure 5 shows a representative
autoradiograph of newly deposited 35S-labeled collagen IV
in RPTC after exposure to DCVC. Densitometric analysis of the 206-kDa
collagen IV band was performed on scanned images of individual
autoradiographs from three separate experiments (Fig.
6). To illustrate changes in collagen IV
deposition over time in each group, values are expressed as a
percentage of day 1 controls grown in 50 µM AscP. There
was a numerical increase in collagen IV deposition by uninjured
RPTC cultured in pharmacological concentrations of AscP compared
with RPTC cultured in physiological concentrations of AscP. However,
this numerical increase was not statistically significant. One day
after DCVC exposure, collagen IV deposition was significantly inhibited
in cells grown in the presence of 50 µM AscP. Collagen IV deposition
was numerically decreased ~50% compared with controls in
DCVC-injured RPTC grown in physiological concentrations of AscP on
day 4 after injury, although this decrease was not
statistically significant. However, DCVC-injured RPTC grown in 500 µM
AscP maintained collagen IV deposition at levels equal to that
of controls throughout the experiment. Compared with
day 1, on days 4 and 6 a decrease in collagen IV deposition was seen in all groups. On day 6,
there were no differences in collagen IV deposition among any
treatment groups. These findings demonstrate that RPTC sublethally
injured by DCVC in the presence of physiological concentrations of
ascorbic acid decrease collagen IV deposition after the injury.
However, in the presence of pharmacological concentrations of ascorbic acid, sublethally injured RPTC maintain collagen IV deposition at
levels equal to controls.
|
|
Proline hydroxylation in collagen IV.
Loss of 3H due to hydroxylation of the 4-trans
position of proline in newly synthesized collagen IV was used as a
marker of prolyl hydroxylase activity. RPTC grown in 50 and 500 µM
AscP were dual labeled with L-[4-3H]proline
and L-[14C]proline over a 24-h period after
DCVC exposure, and the loss of 3H was measured as a
decrease in the ratio of L-[4-3H]proline to
L-[14C]proline. DCVC-injured RPTC grown
in 50 µM AscP had a ratio of L-[4-3H]proline to
L-[14C]proline that was ~80% greater than
controls on day 1 after injury, indicating a
significant decrease in proline hydroxylation (Fig. 7). The
L-[4-3H]proline-to-L-[14C]proline
ratio in DCVC-injured RPTC grown in 500 µM AscP was ~45% greater
than controls on day 1 after injury. These data suggest that
DCVC exposure inhibits prolyl hydroxylase activity and that decreased
proline hydroxylation may contribute to the inhibition of collagen IV
deposition in DCVC-injured RPTC grown in 50 µM AscP. In contrast, the
degree of proline hydroxylation in newly synthesized collagen IV is
higher in DCVC-injured RPTC grown in the presence of 500 µM AscP,
suggesting that ascorbic acid-stimulated prolyl hydroxylase activity
may contribute in part to the deposition of collagen IV in DCVC-injured
RPTC.
|
Effect of exogenous ECM proteins on cell repair.
Because DCVC-injured RPTC cultured in the presence of pharmacological
concentrations of AscP maintain collagen IV deposition and are capable
of physiological repair, we determined whether exogenous renal tubular
ECM proteins promoted cell repair in DCVC-injured RPTC. DCVC-injured
RPTC grown in physiological concentrations of AscP were exposed to
exogenous collagen I or collagen IV (0, 5, 15, 50 µg/ml) immediately
after DCVC exposure and continuously through day 6 after
injury. Basal QO2 and monolayer cell density were measured on days 1 and 6 after injury
induced by DCVC. Exposure of untreated RPTC to collagen I or collagen
IV had no effect on basal QO2 or monolayer cell
density on days 1 or 6 (data not shown). On
day 1 after injury, monolayer cell density and basal
QO2 were decreased ~50% in injured RPTC
grown in the absence or presence of collagen IV (Fig.
8, top, and Fig.
9, top) and collagen I (data not shown). On day 6, cell density in injured RPTC cultured
in the presence of collagen IV remained ~50% of uninjured RPTC and equal to the cell density in DCVC-injured RPTC grown in the absence of
collagen IV (Fig. 8, bottom). However, DCVC-injured RPTC
cultured in the presence of exogenous collagen IV recovered basal
QO2 by day 6 after injury, with a
complete repair in RPTC grown in the presence of 50 µg/ml collagen IV
(Fig. 9, bottom). DCVC-injured RPTC cultured in the presence
of collagen I exhibited neither increased cell density nor repair of
basal QO2 by day 6 after injury
(Fig. 10).
|
|
|
|
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
RPTC that do not die or that become detached from the ECM after ischemic or chemical injury are thought to undergo repair or dedifferentiate, proliferate, and migrate to denuded areas of the tubules. The cells of the newly lined tubule may then differentiate, promoting the return of normal tubular function and overall renal function. During our studies of the mechanisms of RPTC repair and regeneration after toxicant exposure, we observed that RPTC sublethally injured by DCVC neither proliferated nor repaired physiological functions (37). In contrast, RPTC sublethally injured by the oxidant t-butyl hydroperoxide proliferated and repaired physiological functions (35). However, when RPTC sublethally injured by DCVC were cultured in the presence of pharmacological concentrations of AscP, they proliferated and repaired physiological functions (36). Thus one goal of this study was to determine whether one of the mechanisms of action of pharmacological concentrations of ascorbic acid in promoting RPTC repair and regeneration is through the stimulation of the synthesis and deposition of collagen IV.
AscP is known to promote collagen synthesis and deposition in cultured cells (13, 33). Collagen IV is the most abundant component of the proximal tubular BM, and the regulation of collagen IV synthesis and degradation plays an important role in cell function, growth, migration, and organ remodeling in many tissues (15). Furthermore, collagen IV synthesis and deposition are increased in control RPTC exposed to AscP (36). We examined collagen IV synthesis and deposition in RPTC sublethally injured by DCVC. One day after DCVC exposure, collagen IV synthesis increased in RPTC grown in the presence of physiological concentrations of AscP and decreased to control levels on days 4 and 6. In contrast, collagen IV deposition into the ECM was inhibited by DCVC 1 day after exposure, a trend that was still evident on day 4, although the statistical significance of this decrease was not conclusive due to a significant decrease in collagen IV deposition in all treatment groups. These observations represent the novel findings 1) that an increase in collagen IV synthesis is a response to DCVC injury and 2) that one potential toxic mechanism of DCVC-induced nephrotoxicity is the inhibition of collagen IV deposition into the ECM. These observations suggest that the lack of repair and regeneration in DCVC-injured RPTC grown in physiological concentrations of ascorbic acid may result from an inability to deposit collagen IV in response to injury.
Examination of collagen IV synthesis and deposition in RPTC grown in pharmacological concentrations of AscP showed that injury by DCVC increased collagen IV synthesis as was shown in DCVC-injured RPTC grown in physiological concentrations of ascorbic acid. Unlike DCVC-injured RPTC grown in physiological concentrations of AscP, DCVC-injured RPTC grown in pharmacological concentrations of AscP maintained collagen IV deposition at control levels. These data suggest that the maintenance of collagen IV deposition after DCVC exposure promotes RPTC repair and regeneration by day 6 after injury, thus creating an association between the ability of injured RPTC to deposit collagen IV and to repair physiological functions.
One potential explanation for the lack of collagen IV deposition in
DCVC-injured RPTC grown in physiological concentrations of AscP is that
newly synthesized procollagen chains are not being properly
hydroxylated at susceptible proline residues. Underhydroxylated procollagen
chains will not fold into triple helical monomers adequately, and most are targeted for degradation intracellularly rather than being secreted into the ECM (18, 47). Prolyl
hydroxylase is the microsomal enzyme responsible for proline
hydroxylation of procollagen
chains, and ascorbic acid is the
preferred iron-reducing cofactor for prolyl hydroxylase activity
(41). Approximately 45-50% of susceptible collagen
proline residues are hydroxylated in normal vertebrate tissues;
however, when the degree of proline hydroxylation is decreased, so is
the amount of collagen deposited into the ECM (28). In our
study, proline hydroxylation in newly synthesized collagen IV was
decreased after DCVC exposure in RPTC grown in the presence of
physiological concentrations of AscP. However, proline hydroxylation in
injured RPTC grown in pharmacological concentrations of AscP was
greater compared with injured RPTC grown in physiological
concentrations of AscP, suggesting that these cells retain some ability
to hydroxylate susceptible proline residues. Although we have no
evidence of a direct interaction between DCVC and prolyl hydroxylase,
these results suggest that insufficient proline hydroxylation
contributes to decreased collagen IV deposition in DCVC-injured RPTC
grown in physiological concentrations of AscP. In addition,
pharmacological concentrations of AscP increase the degree of proline
hydroxylation in collagen IV after DCVC injury and may promote the
maintenance of collagen IV deposition into the ECM. These results
further support our conclusion that the deposition of collagen IV is
associated with RPTC repair after sublethal injury.
Figure 4 shows that uninjured RPTC grown in pharmacological concentrations of AscP synthesize less collagen IV on day 1 after confluence than RPTC grown in physiological concentrations of AscP. This finding can be explained by the idea that RPTC cultures synthesize less collagen over time, especially after reaching confluence, as the cultures begin to quiesce. This hypothesis is supported by previous studies in regenerating tissues demonstrating upregulated ECM protein expression that returns to basal levels when tissue structure is restored (49). Because ascorbate is known to enhance cellular growth, RPTC grown in pharmacological AscP concentrations may decrease collagen synthesis to basal levels sooner than RPTC grown in physiological concentrations of AscP (39). Further evidence of this effect is observed on day 4 when RPTC cultured in physiological AscP concentrations exhibit collagen IV synthesis that is decreased to levels of RPTC cultured in pharmacological concentrations of AscP.
The next goal of this study was to determine the effect of exogenous collagens I and IV, laminin, and fibronectin on cell repair in RPTC sublethally injured by DCVC. On the basis of the correlation between cell repair and collagen IV deposition, we hypothesized that adding exogenous collagen IV to the culture media would promote cell repair and/or proliferation in DCVC-injured RPTC. In addition, by determining the effects of collagen I, laminin, or fibronectin on cell repair, we would determine whether any effect of collagen IV to stimulate repair is specific to collagen IV or is a general effect of ECM proteins. Collagen IV, but not collagen I, laminin, or fibronectin, promoted repair of mitochondrial oxygen consumption (basal QO2) and active Na+ transport (Na+-K+-ATPase activity) after injury, implicating collagen IV as an important ECM protein involved in the repair of physiological functions in RPTC. However, DCVC-injured RPTC cultured in the presence of collagen IV did not exhibit increased cell density on day 6 after DCVC exposure. This finding suggests that collagen IV is not involved in the proliferation of RPTC as observed in the presence of pharmacological concentrations of AscP and that ascorbic acid plays other important roles in renal recovery not related to ECM production. These data provide strong evidence of a specific role for collagen IV in cell repair, but not proliferation, after sublethal injury.
Despite the prevalence of theories implicating collagen IV as a mediator of renal tubular development and function, the nonpathological role of collagen IV in renal injury, repair, and restoration of tubular function is relatively unknown. The novel findings presented here suggest that extracellular collagen IV specifically promotes the repair of physiological functions in injured RPTC. The ECM proteins collagen I, laminin, and fibronectin did not promote repair of physiological functions. DCVC-injured RPTC exhibited decreased collagen IV hydroxylation and deposition and were unable to repair physiological functions. In contrast, RPTC grown in pharmacological concentrations of AscP maintained collagen IV hydroxylation and deposition and were able to repair physiological functions and increase cell density after DCVC injury. Therefore, there is a correlation between AscP-stimulated hydroxylation and deposition of collagen IV and the ability of injured RPTC to repair physiological functions.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank Dr. Thomas W. Petry (Upjohn Pharmacia, Kalamazoo, MI) for his generous gift of DCVC.
![]() |
FOOTNOTES |
---|
This work was supported by National Institute of Environmental Health Sciences Grant ES-O4410 (R. G. Schnellmann) and a predoctoral fellowship from the American Heart Association, Heartland Affiliate (P. A. Nony).
Address for reprint requests and other correspondence: R. G. Schnellmann, Dept. of Pharmaceutical Sciences, 280 Calhoun St., POB 250140, Charleston, SC 29425.
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. Section 1734 solely to indicate this fact.
Received 28 July 2000; accepted in final form 20 April 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Abbate, M,
and
Remuzzi G.
Acceleration of recovery in acute renal failure: from cellular mechanisms of tubular repair to innovative targeted therapies.
Ren Fail
18:
377-388,
1996[ISI][Medline].
2.
Alcain, FJ,
and
Buron MI.
Ascorbate on cell growth and differentiation.
J Bioenerg Biomembr
26:
393-398,
1994[ISI][Medline].
3.
Bailey, AJ,
Sims TJ,
Duance VC,
and
Light ND.
Partial characterization of a second basement membrane collagen in human placenta: evidence for the existence of two type IV collagen molecules.
FEBS Lett
99:
361-366,
1979[ISI][Medline].
4.
Cantley, LG.
Growth factors and the kidney: regulation of epithelial cell movement and morphogenesis.
Am J Physiol Renal Fluid Electrolyte Physiol
271:
F1103-F1113,
1996
5.
Chojkier Peterkofsky, MB,
and
Bateman J.
A new method for determining the extent of proline hydroxylation by measuring changes in the ratio of [4-3H]:[14C]proline in collagenase digests.
Anal Biochem
108:
385-393,
1980[ISI][Medline].
6.
Davidson, JM,
LuValle PA,
Zoia O,
Quaglino D, Jr,
and
Giro M.
Ascorbate differentially regulates elastin and collagen biosynthesis in vascular smooth muscle cells and skin fibroblasts by pretranslational mechanisms.
J Biol Chem
272:
345-352,
1997
7.
Denk, PO,
and
Knorr M.
In vitro effect of ascorbic acid on the proliferation of bovine scleral and Tenon's capsule fibroblasts.
Eur J Ophthalmol
8:
37-41,
1998[ISI][Medline].
8.
Dixit, SN.
Isolation, purification and characterization of intact and pepsin-derived fragments of laminin from human placenta.
Connect Tissue Res
14:
31-40,
1985[ISI][Medline].
9.
Englard, S,
and
Seifter S.
The biochemical functions of ascorbic acid.
Annu Rev Nutr
6:
365-406,
1986[ISI][Medline].
10.
Geesin, JC,
Darr D,
Kaufman R,
Murad S,
and
Pinnell SR.
Ascorbic acid specifically increases type I and type III procollagen messenger RNA levels in human skin fibroblasts.
J Invest Dermatol
90:
420-424,
1988[Abstract].
11.
Ghohestani, RF,
Hudson BG,
Claudy A,
and
Uitto J.
The 5 chain of type IV collagen is the target of IgG autoantibodies in a novel autoimmune disease with subepidermal blisters and renal insufficiency.
J Biol Chem
275:
16002-16006,
2000
12.
Gibbs, SR,
Goins RA,
Belvin EL,
Dimari SJ,
Merriam AP,
Bowling-Brown S,
Harris RC,
and
Haralson MA.
Characterization of the collagen phenotype of rabbit proximal tubule cells in culture.
Connect Tissue Res
40:
173-188,
1999[ISI][Medline].
13.
Hata, RI,
and
Senoo H.
L-Ascorbic acid 2-phosphate stimulates collagen accumulation, cell proliferation, and formation of a three-dimensional tissue-like substance by skin fibroblasts.
J Cell Physiol
138:
8-16,
1989[ISI][Medline].
14.
Houglum, KP,
Brenner DA,
and
Chojkier M.
Ascorbic acid stimulation of collagen biosynthesis independent of hydroxylation.
Am J Clin Nutr
54:
1141S-1143S,
1991[Abstract].
15.
Hudson, BG,
Reeders ST,
and
Tryggvason K.
Type IV collagen: structure, gene organization, and role in human diseases: molecular basis of Goodpasture and Alport syndromes and diffuse leiomyomatosis.
J Biol Chem
268:
26033-26036,
1993
16.
Ivanov, VO,
Rabovsky AB,
Ivanova SV,
and
Niedzwiecki A.
Transforming growth factor-beta 1 and ascorbate regulate proliferation of cultured smooth muscle cells by independent mechanisms.
Atherosclerosis
140:
24-25,
1998.
17.
Juva, K,
Prockop DJ,
Cooper G,
and
Lash J.
Hydroxylation of proline and the intracellular accumulation of a polypeptide precursor of collagen.
Science
152:
92-94,
1966[ISI][Medline].
18.
Kalluri, R,
and
Cosgrove D.
Assembly of type IV collagen: insights from 3(IV) collagen-deficient mice.
J Biol Chem
275:
12719-12724,
2000
19.
Kalluri, R,
Shield CF, III,
Todd P,
Hudson BG,
and
Neilson EG.
Isoform switching of type IV collagen is developmentally arrested in X-linked Alport syndrome leading to increased susceptibility of renal basement membranes to endoproteolysis.
J Clin Invest
99:
2470-2478,
1997
20.
Kao, WW,
Prockop DJ,
and
Berg RA.
Kinetics for the secretion of non-helical procollagen by freshly isolated tendon cells.
J Biol Chem
254:
2234-2243,
1979[Abstract].
21.
Komsa-Penkova, R,
Koynova R,
Kostov G,
and
Tenchov BG.
Thermal stability of calf skin collagen type I in salt solutions.
Biochim Biophys Acta
1297:
171-181,
1996[ISI][Medline].
22.
Laemmli, UK.
Cleavage of structural proteins during the assembly of the head bacteriophage T4.
Nature
227:
680-685,
1970[ISI][Medline].
23.
Lash, LH,
Sausen PJ,
Duescher RJ,
Cooley AJ,
and
Elfarra AA.
Roles of cysteine conjugate -lyase and S-oxidase in nephrotoxicity: studies with S-(1,2-dichlorovinyl)-L-cysteine and S-(1,2-dichlorovinyl)-L-cysteine sulfoxide.
J Pharmacol Exp Ther
269:
374-383,
1994[Abstract].
24.
Levine, M.
New concepts in the biology and biochemistry of ascorbic acid.
N Engl J Med
314:
892-902,
1986[ISI][Medline].
25.
Linsenmayer, TF.
Cell Biology of the Extracellular Matrix. New York: Plenum, 1991, p. 5-37.
26.
Lowry, OH,
Rosebrough NJ,
Farr AL,
and
Randall RJ.
Protein measurement with the Folin phenol reagent.
J Biol Chem
193:
265-275,
1951
27.
Martensson, S,
Brunmark C,
Ohlsson L,
Bak-Jensen E,
Butkowski R,
Boketoft A,
and
Wieslander J.
Heterogeneity of renal cell carcinoma.
Nephrol Dial Transplant
10:
1637-1643,
1995[Abstract].
28.
Miller, EJ.
Isolation and characterization of a collagen from chick cartilage containing three identical alpha chains.
Biochem J
10:
1652-1659,
1971.
29.
Miner, JH.
Renal basement membrane components.
Kidney Int
56:
2016-2024,
1999[ISI][Medline].
30.
Moore, RB,
and
Green T.
The synthesis of nephrotoxin conjugates of glutathione and cysteine.
Toxicol Environ Chem
17:
153-162,
1988[ISI].
31.
Müller, U,
and
Brändli AW.
Cell adhesion molecules and extracellular-matrix constituents in kidney development and disease.
J Cell Sci
112:
3855-3867,
1999
32.
Murad, S,
Grove D,
Lindberg KA,
Reynolds G,
Sivarajah A,
and
Pinnell SR.
Regulation of collagen synthesis by ascorbic acid.
Proc Natl Acad Sci USA
78:
2879-2882,
1981[Abstract].
33.
Murad, S,
Tajime S,
Johnson GR,
Arunthathy S,
and
Pinnell SR.
Collagen synthesis in cultured human skin fibroblasts: effect of ascorbic acid and its analogs.
J Invest Dermatol
81:
158-162,
1983[Abstract].
34.
Niki, T,
Schuppan D,
De Bleser PJ,
Vrijsen R,
Pipeleers-Marichal M,
Beyaert R,
Wisse E,
and
Geerts A.
Dexamethasone alters messenger RNA levels but not synthesis of collagens, fibronectin, or laminin by cultured rat fat-storing cells.
Hepatology
23:
1673-1681,
1996[ISI][Medline].
35.
Nowak, G,
Aleo MD,
Morgan JA,
and
Schnellmann RG.
Recovery of cellular functions following oxidant injury.
Am J Physiol Renal Physiol
274:
F509-F515,
1998
36.
Nowak, G,
Carter CA,
and
Schnellmann RG.
Ascorbic acid promotes recovery of cellular functions following toxicant-induced injury.
Toxicol Appl Pharmacol
167:
37-45,
2000[ISI][Medline].
37.
Nowak, G,
Keasler KB,
McKeller DE,
and
Schnellmann RG.
Differential effects of EGF on repair of cellular functions after dichlorovinyl-L-cysteine-induced injury.
Am J Physiol Renal Physiol
276:
F228-F236,
1999
38.
Nowak, G,
and
Schnellmann RG.
Integrative effects of EGF on metabolism and proliferation in renal proximal tubular cells.
Am J Physiol Cell Physiol
269:
C1317-C1325,
1995
39.
Nowak, G,
and
Schnellmann RG.
L-Ascorbic acid regulates growth and metabolism of renal cells: improvements in cell culture.
Am J Physiol Cell Physiol
271:
C2072-C2080,
1996
40.
Peterkofsky, B.
Ascorbate requirement for hydroxylation and secretion of procollagen: relationship to inhibition of collagen synthesis in scurvy.
Am J Clin Nutr
54:
1135S-1140S,
1991[Abstract].
41.
Pinnell, S.
Regulation of collagen biosynthesis by ascorbic acid: a review.
Yale J Biol Med
58:
553-559,
1985[ISI][Medline].
42.
Rosenbloom, J,
and
Prockop DJ.
Incorporation of 3,4-dehydroproline into protocollagen and collagen. Limited hydroxylation of proline and lysine in the same polypeptide.
J Biol Chem
245:
3361-3368,
1970
43.
Schmidt, C,
Pollner R,
Pöschl E,
and
Kühn K.
Expression of human collagen type IV genes is regulated by transcriptional and post-transcriptional mechanisms.
FEBS Lett
312:
174-178,
1992[ISI][Medline].
44.
Schwartz, GJ,
and
Evan AP.
Development of solute transport in rabbit proximal tubule. III. Na-K-ATPase activity.
Am J Physiol Renal Fluid Electrolyte Physiol
246:
F845-F852,
1984[ISI][Medline].
45.
Shapiro, IM,
Leboy PS,
Tokuoka T,
Forbes E,
DeBolt K,
Adams SL,
and
Pacifici M.
Ascorbic acid regulates multiple metabolic activities of cartilage cells.
Am J Clin Nutr
54:
1209S-1213S,
1991[Abstract].
46.
Sullivan, TA,
Uschmann B,
Hough R,
and
Leboy PS.
Ascorbate modulation of chondrocyte gene expression is independent of its role in collagen excretion.
J Biol Chem
269:
22500-22506,
1994
47.
Tschank, G,
Hanauske-Abel HM,
and
Peterkofsky B.
The effectiveness of inhibitors of soluble prolyl hydroxylase against the enzyme in cisternae of isolated bone microsomes.
Arch Biochem Biophys
261:
312-323,
1988[ISI][Medline].
48.
Venezian, R,
Shenker BJ,
Datar S,
and
Leboy PS.
Modulation of chondrocyte proliferation by ascorbic acid and BMP-2.
J Cell Physiol
174:
331-341,
1996.
49.
Walker, PD.
Alterations in renal tubular extracellular matrix components after ischemia-reperfusion injury to the kidney.
Lab Invest
70:
229-345,
1994.
50.
Wewer, U,
Albrechtsen R,
Manthorpe M,
Varon S,
Engvall E,
and
Ruoslahti E.
Human laminin isolated in a nearly intact, biologically active form from placenta by limited proteolysis.
J Biol Chem
258:
12654-12660,
1983
51.
Yamada, KM,
and
Kennedy DW.
Fibroblast cellular and plasma fibronectins are similar but not identical.
J Cell Biol
80:
492-498,
1979[Abstract].