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Regular Article |
From the Department of Medicine and Hypertension,*
Division of Nephrology and Hypertension, and the Department of
Pathology and Laboratory Medicine,
University
of North Carolina at Chapel Hill, Chapel Hill, North Carolina; and the
Department of Nephrology,
Lund University,
Lund, Sweden
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Structurally, PR3 belongs to the serprocidin subgroup
of the chymotrypsin-like protease superfamily, comprised of neutrophil
elastase, cathepsin G, and azurocidin.15-17
With the
exception of azurocidin, which is a pseudoprotease, these proteases
degrade extracellular matrix macromolecules, such as elastin,
fibronectin, laminin, vitronectin, and type IV
collagen.4,5,18,19
Sequence analysis of PR3 showed it to
be identical to myeloblastin, a gene cloned as a differentiation marker
from promyelocytic leukemia HL60 cells.20
PR3 has been
shown to be involved in retinoic acid-induced differentiation of HL60
cells.20
Apparently, retinoic acid indirectly activates
PR3, which then hydrolyzes hsp27/28, a component of mitogenic signal
transduction pathways.21,22
Other known substrates of PR3
include transcription factors (nuclear factor-
B and SP1) and
cytokines (tumor necrosis factor-
, transforming growth factor-{beta}1,
and interleukin-1{beta}).23-26
Although proteolytic activity
of PR3 is involved in processing these molecules, there are
accumulating data that indicate that PR3 can have direct effects on
intracellular processes in the absence of proteolytic activity. For
example, a secreted, inactive proform of PR3 is shown to down-modulate
DNA synthesis in normal hematopoietic progenitor cells. The inhibitory
effect of secreted PR3 is reversible by granulocyte-macrophage
colony-stimulating factor, implying that PR3 can function as a
counterbalance to regulators of proliferation.27
Enzymatically inactive PR3 has been shown to induce interleukin-8
production, whereas enzymatic activity is essential for elastase to
stimulate this effect.28
We have reported that inactive
PR3 induced apoptosis of bovine pulmonary artery endothelial
cells29
and subsequent studies confirmed these
findings.30
Hence, PR3 seems to be a multifunctional
protein endowed with the capacity to influence cell cycle,
differentiation, and cell death.
It has been suggested that pulmonary injury and renal glomerular damage may be caused by the MPO system. MPO degrades H2O2 in the presence of chloride to produce hypochlorite (OCl-). The products of the MPO-H2O2-chloride system are powerful oxidants that can have profound biological effects. When MPO and H2O2 are released to the outside of the cell, a reaction with chloride can induce damage to adjacent tissue and thus contribute to the pathogenesis of disease.13,14
The important issue addressed by the study presented here is the mechanism of neutrophil-mediated damage to endothelial and epithelial cells during inflammation. We investigate whether or not granule proteins, PR3 and MPO, are internalized into endothelial and epithelial cells and whether internalization has an effect on these cells. Precedence for nuclear import of granule proteases is provided by reports showing that granzyme B, a protease from granules of T lymphocytes and natural killer cells and a homologue of PR3, can be transported to the nucleus in treated Cos cells and the cells die by apoptosis.31 Signal transduction events, associated with granzyme B-induced apoptosis, included activation of the cyclin-dependent kinase cdc-2.32 Further, it was shown that specific inhibitors of the proteolytic activity of granzyme B had no effect on the nuclear import, implying that proteolytic activity was not essential for nuclear targeting.
We report that human endothelial cells bind, sequester, and route neutrophil proteins, PR3, and MPO, from the cell surface to the nucleus where they co-localize with chromatin. PR3 internalization is concomitant with apoptosis in contrast to MPO. Epithelial cells do not have the capacity to internalize PR3 and do not die by apoptosis. The apoptotic function of PR3 was mapped to a 100-amino acid fragment of the molecule, which contains no component of the catalytic triad. MPO internalization is not restricted by cell type, implying that the mechanism is not the same as that for PR3 internalization. Internalized MPO confers increased intracellular oxidant production.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Granule proteins were isolated from leukocytes of human donors with leukemia.29 PR3 was purified by a monoclonal anti-PR3 affinity column (monoclonal anti-PR3 IgG, 12-1D6-1D4, developed in our laboratory) and a Bio-Rex 70 column (Bio-Rad Laboratories, Richmond, CA) and MPO by a Resource S cation exchange column (Pharmacia, Uppsala, Sweden) and a Superdex 200 gel filtration column (Pharmacia).29 The purity of PR3 and MPO was determined by amino acid sequencing, enzyme-linked immunosorbent assay, Western blot analysis, and enzymatic activity. Enzymatic activity of MPO was 172 U/mg protein using 4-aminoanipyrine as substrate.33 PR3 proteolytic activity was very low, using Boc-Ala-Onp as substrate.29 Protein preparations were determined to be endotoxin-free. Proteolytically active PR3 (109 U/mg protein) was a generous gift from Dr. J. Wieslander (Wieslab AB, Lund, Sweden). Triton X-100 was removed from active PR3 using Extracti-Gel D AffinityPak detergent-removing column (Pierce, Rockford, IL).
Cell Culture Conditions
Pooled human umbilical vein endothelial cells (HUVECs), single-donor human umbilical arterial endothelial cells (HUAECs), single-donor human lung microvascular endothelial cells (HMVEC-Ls), and single-donor human small airway epithelial cells (SAECs) were obtained from Clonetics (San Diego, California, CA). HUVECs and HUAECs were cultured in EGM BulletKit, HMVEC-Ls in EGM-2 MV BulletKit, and SAECs in SAGM BulletKit media (Clonetics). Media contained growth factors, 2% fetal calf serum (FCS), antibiotics, and other supplements, but there was no FCS in the SAGM BulletKit medium. HUVECs and SAECs, passage 3 or 4, and HUAECs and HMVEC-Ls, passage 5 or 6, were used for experiments. Monkey epithelial cells (Cos-7) were maintained in Dulbecco’s modified Eagle’s medium-H, 10% fetal bovine serum, and antibiotics. EA.hy926 cells (HUVECs fused to A549 human lung epithelial carcinoma) were cultured in the same medium plus hyproxanthine aminopterin thymidine (HAT) supplement.
Translocation of Granule Proteins into Cells
Cells on microscope cover glasses were incubated with 10 µg/ml of PR3 or MPO for 5 minutes to 16 hours at 37°C in media without growth factors or FCS. For immunofluorescent staining, cells were fixed with 2% paraformaldehyde in phosphate-buffered saline (PBS), permeabilized with acetone, and blocked with 0.2 mol/L glycine and 5% donkey serum in PBS. Primary antibodies were rabbit anti-PR3 (Wieslab AB) or rabbit anti-MPO antibody (DAKO, Carpinteria, CA) and secondary antibody, fluorescein isothiocyanate-conjugated affinipure F(ab')2 fragment donkey anti-rabbit IgG (H+L) (Jackson, West Grove, PA), was used. Cells were mounted and viewed by a Nikon FXA research light microscope (Nikon, Garden City, NY) for standard immunofluorescence staining. A Zeiss confocal laser-scanning microscope LSM 110 (Carl Zeiss, Thornwood, NY) for confocal immunofluorescence staining, with an optical section thickness is 0.5 µm using the 63x plan apo 1.4 NA objective. The cells were analyzed from top to bottom in 15 different planes. The appropriate negative controls were performed concurrently, consisting of untreated cells stained with primary antibodies and/or secondary antibody, PR3-treated cells with anti-MPO antibody as primary antibody and MPO-treated cells with anti-PR3 antibody as primary antibody, and treated cells with primary antibodies or secondary antibody alone.
For immunogold labeling, cells were processed according to the methods described by Madden.34 Samples were blocked in 5% goat serum, incubated with primary antibodies, monoclonal anti-PR3 IgG (4A3) (Wieslab AB) or polyclonal anti-MPO IgG (DAKO), for 2 hours, and subsequently incubated with secondary antibodies, goat anti-mouse 10-nm colloidal gold (Amersham Life Science, Arlington Heights, IL) for monoclonal primary antibody or protein A 10 nm colloidal gold (Ted Pella, Redding, CA) for polyclonal primary antibody, for 1 hour at room temperature. The grids were observed and photographed using a LEO EM-910 transmission electron microscopy (LEO Electron Microscopy, Thornwood, NY). The appropriate negative controls were performed concurrently, consisting of untreated cells stained with anti-PR3 IgG or anti-MPO IgG and treated cells stained with normal mouse or rabbit IgG (DAKO).
Assessment of Apoptosis
Cells in flasks were treated with 1 to 10 µg/ml of isolated and enzymatically inactive PR3 or 1 to 100 µg/ml MPO for 6 to 24 hours in medium containing 2% FCS, but without growth factors. For all assays, both detached and adhered cells were harvested and combined. Apoptosis was assessed by characteristic morphological changes using an UV light microscope (Nikon), and by DNA content using flow cytometry. For UV light microscopy, cells were stained with DNA dyes, either 1 µg/ml 4',6-diamidino-2-phenylindole (DAPI) (Sigma, St. Louis, MO) or 20 µg/ml propidium iodide (Sigma). Apoptotic cells were identified by chromatin condensation and fragmentation.35 Data reported was from six samples of at least three independent experiments, counting 500 cells from randomly selected fields. For flow cytometry, cells were fixed in 70% ethanol/PBS, incubated at 4°C overnight, and resuspended in propidium iodide staining mixture, as described previously.29 DNA content was determined with a FACScan flow cytometer (Becton Dickinson Immunocytometry Systems, San Jose, CA) linked to a Cicero/Cyclops system (Cytomation, Fort Collins, CO) for data acquisition and analysis. Apoptotic cells were identified by a decrease in DNA content resulting in an apoptotic subG0/G1 population peak. At least 20,000 cells were analyzed in each sample and data were reported from at least three independent experiments.
Subcloning of PR3-N, PR3-M, and PR3-C cDNA Sequences
PR3cDNA/pAcC4 plasmid was a generous gift from Dr. Joelle E. Gabay, Cornell University Medical College. Nucleotides 1–313 (PR3-N), 314–604 (PR3-M), and 620–741 (PR3-C) were amplified by polymerase chain reaction (PCR). Primers were designed to generate restriction sites at each end and a kozak sequence at the 5' end. Parameters were 30 cycles of 94°C for 30 seconds, 62°C for 30 seconds, and 72°C for 2 minutes. The primers for PR3-N cDNA were 5'-GATATCGCCACCATGGGCGCTCACCGGCCCC-3' (forward) and 5'-TGGAATTCTCTGAGCCACCGAGAAGTG-3' (reverse); PR3-M cDNA were 5'-GATATCGCCACCATGGGCGTGTTTCTGAACAACTACGAC-3' (forward) and 5'-TGGAATTCTTCCGAAGCAGATGCCGGCCTT-3' (reverse), and PR3-C cDNA primers were 5'-GATATCGCCACCATGGGCCTGATCTGTGATGGCATCATC-3' (forward) and 5'-TGGAATTCTCAGCGTGGAACGGATCCAGT-3' (reverse). Amplification products were subcloned into pCR-Blunt (Zero Blunt PCR Cloning Kit; Invitrogen, Carlsbad, CA) and sequenced using an M13 forward primer (5'-GTAAAACGACGGCCAG-3') (UNC-CH Automated DNA Sequencing Facility, Chapel Hill, NC). Insert was retrieved from pCR-blunt and subcloned into pcDNA3.1(-)/Myc-HisA (Invitrogen)
Analysis of Apoptosis Induced by PR3 Fragments
Cos-7 cells were electroporated 24 hours after trypsinization with 30 µg of plasmid DNA in serum-free RPMI at 300 V and 960 µF using the Gene Pulser II Electroporation System (Biorad, Hercules, CA). Cells were cultured for 48 hours and conditioned medium was collected, centrifuged, and added to EA.hy926 cells. Apoptosis was quantitated by FACScan analysis of terminal dUTP nick-end labeling (TUNEL) cells using fluorescein isothiocyanate-dUTP and terminal transferase (TdT) (Boehringer Mannheim, Mannheim, Germany). Briefly, the cells were fixed with 1% paraformaldehyde in PBS and permeabilized with 70% ethanol and labeling reaction was performed. One half of each cell sample was used to determine background fluorescence by omitting TdT from the reaction mixture. The TdT was added to the second half and increases in fluorescence (FL1) were evaluated. Clumps and doublets were excluded by using forward scatter versus propidium iodide fluorescence (FL2-W).
Detection of Intracellular Oxidants Generated by MPO
Intracellular oxidants generation was detected by oxidation of nonfluorescent dihydrorhodamine (DHR) (New Concept Scientific. Burlington, Ontario, Canada) to fluorescent rhodamine. Cells were incubated with 10 µmol/L DHR for 1 hour in medium without FCS or growth factors, with the subsequent addition of MPO (1 to 50 µg/ml) or PR3 (1 to 10 µg/ml) at 37°C for 4 to 24 hours. Intracellular oxidant generation was determined by three methods. Cell fluorescence was observed by UV light microscopy. By flow cytometry, cells were harvested with trypsin/ethylenediaminetetraacetic acid and cell fluorescence were examined in FL1 green channel in which at least 20,000 cells were analyzed. For fluorometry, treated cells were washed, harvested, and lysed by sonication in 20 mmol/L Tris-HCl, pH 7.4, buffer. The fluorescence of cell lysates was measured by a fluorometer FLUOstar 403 (BMG Lab Technologies, Offenburg, Germany), 5 time flashes per sample (excitation wavelength of 485 nm, emission wavelength of 538 nm).36 For blocking studies, MPO was pre-incubated with a 10-fold dose of catalase (Calbiochem, La Jolla, CA) for 30 minutes before the addition to cells. The appropriate negative controls were performed concurrently, consisting of cells incubated with medium or DHR alone, and no cell plate incubated with DHR and granule proteins.
Statistical Analysis
Analysis of variance was used to determine whether any differences between group means were seen at specific dose and time points within each experiment. When overall differences were found within the experiment, Dunnett’s t-test was used to further compare means of each treatment and its associated dose to the mean of the control group specific to the experiment.37 This test evaluates the minimum significant difference between each group and control as compared to the critical value of Dunnett’s T at the P value of 0.05, whereas controlling for multiple testing. PROC ANOVA and PROC GLM in SAS were used for calculating the statistical analyses (SAS/STAS User’s Guide, SAS Institute, Cary, NC, 1989).
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Results |
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Translocation of PR3 into Endothelial Cells
To examine potential mechanisms of PR3-induced apoptosis, we
determined whether endothelial cells had the capacity to internalize
PR3. PR3-treated cells were examined by three standard techniques:
immunofluorescence microscopy, immunofluorescence confocal laser
scanning microscopy, and electron microscopy. By immunofluorescence
microscopy, HUVECs treated with inactive PR3 showed surface and
intracellular staining by 10 minutes (Figure 1B)
. HUVECs that had not been exposed to
PR3 did not show staining (Figure 1A
and Figure 2A
). After 2 hours, the intensity of the
fluorescence increased, indicating that the level of PR3 in the
cytoplasm of the cell had increased (Figure 1C)
. To confirm these data
and to address the issue of nuclear localization, the PR3-treated cells
were analyzed using confocal laser scanning microscopy. Selected
micrographs are shown in Figure 2
. PR3 staining was detected on the
cell surface, with diffuse staining in both the cytoplasm and the
nucleus by 10 minutes (Figure 2B)
. By 2 hours, the level of PR3
staining increased, indicating that PR3 uptake continued throughout the
2 hours of incubation (Figure 2C)
.
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. After 10 minutes of treatment,
PR3 was detected in cytoplasmic extrusions (Figure 3A)
. We also
observed PR3 on the surface membrane, at attachment plaques, in the
nucleus and in the nucleoli (data not shown). By 2 hours, PR3 was
localized in the nucleus and associated with nuclear heterochromatin
(Figure 3C)
, in nucleoli, in the cytoplasm, and in secondary lysosomes,
but not on the cell surface or in attachment plaques (data
not shown). Endothelial cells harvested from artery (HUAECs) and
lung microvascular (HMVEC-Ls), when incubated with PR3, showed staining
patterns similar to those described above for HUVECs (data not shown).
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We determined if endothelial cells internalize MPO. Analysis of
MPO-treated cells showed that MPO was translocated into HUVECs with a
staining pattern similar to that of PR3. Examination of HUVECs treated
with MPO for 10 minutes showed weak intracellular staining, as assessed
by immunofluorescence microscopy (Figure 1F)
. After 2 hours, strong
cytoplasmic staining was observed displaying a punctate pattern (Figure 1G)
. Analysis by confocal laser scanning microscopy showed prominent
surface membrane staining of MPO by 10 minutes (Figure 2F)
, with some
staining in the cytoplasm and nucleus, and by 2 hours a strong
cytoplasmic signal was detected with some nuclear staining (Figure 2G)
.
By electron microscopy, MPO was localized with heterochromatin in the
nucleus within 10 minutes (Figure 3B)
. Additional staining was observed
at the plasma membrane, in the cytoplasm, at attachment plaques, in the
nucleus and in the nucleoli (data not shown). Within 2 hours, MPO could
be localized to secondary lysosomes (Figure 3D)
, in the cytoplasm,
nucleus and nucleoli, but not on the cell surface and not in attachment
plaques (data not shown). Endothelial cells harvested from artery
(HUAECs) and lung microvascular (HMVEC-Ls), when incubated with MPO,
showed staining patterns similar to those described above for HUVECs
(data not shown).
Translocation of MPO, but Not PR3, into Epithelial Cells
To determine whether translocation of PR3 and MPO was a phenomenon
specific to endothelial cells, or if other cell types could internalize
these proteins, epithelial cells (SAECs) were treated and examined as
described above. Internalization of PR3 could not be detected at any
time point, with very weak surface staining at 2 hours (Figure 1D)
, and
similar results were obtained by confocal microscopy and electron
microscopy (Figure 2D)
. Surprisingly, SAECs, treated with PR3, began to
float off of the plate at 2 hours (
30%) to 4 hours (60%). We
analyzed the cells at multiple time points before detachment (5
minutes, 10 minutes, 15 minutes, 30 minutes, 1 hour, and 2 hours), and
after detachment (4 hours) for PR3 staining. Of cells that were still
attached at 2 hours, no PR3 internalization was detected. We reasoned
that, even if the SAECs had fewer receptors for PR3, some positivity
should have been detected by this time.
In contrast, MPO was internalized by SAECs and by 2 hours prominent
staining was localized around the nucleus (Figure 1H)
. Confocal
microscopy showed similar staining (Figure 2H)
. Analyses of
internalization by electron microscopy showed staining patterns similar
to those described above for HUVECs, with the exception that MPO was
not found in the nucleus in SAECs (data not shown). These data imply
that MPO may enter SAECs through a mechanism different from that of
PR3.
These studies indicate that neutrophil granule proteins, MPO and PR3, cross the endothelial cell membrane. The electron microscopy studies indicate that both proteins can localize to the nucleus. We report that this phenomenon is not unique to endothelial cells harvested from a specific site, but is a common feature observed in HUVECs, HUAECs, and HMVEC-Ls. Interestingly, lung small airway epithelial cells do not seem to have the capability to take up PR3, but are highly receptive to MPO internalization, strongly suggesting that the mechanisms of internalization are unique.
Association of Apoptosis with Internalization of PR3, but Not MPO
To determine whether there is an association between
internalization of PR3 and PR3-induced apoptosis, endothelial cells
were treated with inactive PR3 and apoptosis was quantified. Flow
cytometry was used to detect cells with a less than
G0/G1 DNA content,
indicative of apoptosis, displayed as a
subG0/G1 peak (Figure 4A)
. A dose- and time-dependent increase
in apoptosis was found in PR3-treated HUVECs, compared to controls:
19.9 ± 6.7% versus 7.6 ± 1.7% with 10 µg/ml
for 12 hours; 16.0 ± 3.7% versus 8.0 ± 3.0%
with 5 µg/ml for 24 hours; and 28.4 ± 1.1% versus
8.0 ± 3.0% with 10 µg/ml for 24 hours.
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. HUVECs treated with PR3
demonstrated fragmented nuclei of different sizes (Figure 4C)
. A
statistically significant increase in apoptotic HUVECs was observed in
a dose- and time-dependent manner with 11.7 ± 2.8% apoptotic
cells in 10 µg/ml of PR3 for 24h (Figure 4, D and E)
. To determine
whether inactive PR3 would induce apoptosis in endothelial cells
harvested from sites other than umbilical vein, cells from umbilical
artery and lung microvasculature were analyzed for
fragmented, condensed DNA. Arterial endothelial cells,
HUAECs, treated with 10 µg/ml of PR3 for 24 hours showed 10.3 ±
3.1% apoptosis, whereas lung microvascular cells, HMVEC-Ls, showed
only 3.0 ± 1.8%. Compared with control values of 1.5 ±
1.1% in HUVECs, 1.3 ± 0.8% in HUAECs and 1.0 ± 0.9% in
HMVEC-Ls, the changes were statistically significant with PR3 treatment
in HUVECs and HUAECs, but not in HMVEC-Ls.
These results provide evidence that PR3-induced apoptosis may be linked
with internalization of the protein. If this hypothesis is correct,
then SAECs, which did not internalize PR3, should not undergo apoptosis
with PR3 treatment. Examination of PR3-treated SAECs showed no
morphological changes, by UV light microscopy, indicative of apoptosis,
nor were any apoptotic cells detected by flow cytometry (0.69% with 10
µg/ml of PR3 for 7 hours; controls 0.68%). Of note, SAECs treated
with inactive PR3 began to detach from the dish by 2 hours and a
majority of the cells were floating by 7 hours (90%). Both attached
and detached cells were harvested analyzed and no apoptosis was
detected.
We determined whether internalization of MPO by HUVECs had an
apoptosis-inducing effect. Assessment of nuclear fragmentation by UV
light microscopy showed that MPO did not trigger apoptosis, even at
doses up to 100 µg/ml and for a time up to 24 hours (Figure 4, D and E)
. Nor were any apoptotic cells (above background) detected by flow
cytometry, including cells treated with MPO concentrations ranging from
1 to 100 µg/ml for 6 to 24 hours. Additionally, no apoptosis was
detected in HUAECs, HMVEC-Ls and SAECs after MPO internalization (data
not shown).
These studies strongly suggest that PR3-induced apoptosis requires internalization. The mere process of internalization of extracellular proteins alone is not sufficient to cause apoptosis, based on data showing that MPO internalization does not result in apoptosis.
Identification of a Noncatalytic Domain of PR3 with Apoptotic Function
We have proposed that the apoptosis-inducing effects of PR3
described above were independent of proteolytic activity. The potential
exists that the PR3 preparations used in those studies had very low
levels of proteolytic activity. We hypothesized that if the proteolytic
activity of PR3 is solely responsible for the apoptotic effects on
HUVECs, then blocking this activity should block apoptosis. To test
this, proteolytically active PR3 was used to induce apoptosis in
HUVECs, plus and minus inhibitors. We found that addition of 2% fetal
calf serum blocked only 35.7% apoptosis and addition of
-1-antitrypsin (1-AT) blocked only 36.1%. Because not all
apoptosis was blocked, the data support the hypothesis that PR3 can
induce apoptosis through a mechanism (s) that is independent of its
proteolytic function. However, there is the possibility the protease
inactivators were only partially effective in blocking PR3 activity.
Perhaps they are effective in vitro but once the proteinases
enter the cell, inhibitor activity is lost. To address this issue, we
developed a system to produce PR3 fragments that did not contain the
full component of sequences required for the catalytic site. Based on
crystal structure studies (Figure 5A)
,
only one of three components of the catalytic triad was in the first
one-third of the molecule, the second component was in the middle
portion and the third component was in the last third of the molecule.
The catalytic triad forms the active site through protein
folding.38
Therefore, three unique fragments of PR3 were
generated, each of which contained only one portion of the catalytic
triad. These were subcloned into a mammalian expression vector that
contains a myc-epitope tag and a histidine tail (pcDNA3.1 myc-his).
Cos-7 cells were transiently transfected with vectors coding for the
N-terminal peptide (amino acids 2–104), middle peptide (amino acids
105–204), or the C-terminal peptide (amino acids 205–247) and stained
with a DNA dye to detect DNA fragmentation. Cos-7 cells transfected
with the C-terminal peptide died by apoptosis 24 hours after
transfection, showing fragmented nuclei (Figure 5B)
. Cells transfected
with PR3-N or PR3-M did not show this morphology (data not shown).
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. The terminal transferase enzyme (TdT)
required for end-labeling of the fragmented DNA was omitted to set
parameters of background fluorescence (Figure 5C)
. Increases because of
addition of TdT indicate TUNEL-labeled apoptotic cells. EA.hy926
incubated with conditioned medium (24 hours) from PR3-C-transfected
cells had 20 ± 3.0% TUNEL-positive cells (Figure 5D)
.
PR3-M-transfected cells had 4.75 ± 0.75% positive cells and
PR3-N-transfected cells had 1.07 ± 2.6%. The data indicate that
the C-terminal domain of PR3 can function as a pro-apoptotic molecule,
at least in the cell types tested, whereas the N-terminal, and middle
fragments cannot. These findings support the hypothesis that the
mechanism of PR3-induced apoptosis does not necessarily require its
proteolytic function. Association of Increased Intracellular Oxidation with Internalization of MPO, but Not PR3
The data presented above indicate that internalization of PR3 can
induce apoptosis in endothelial cells, but internalization of MPO does
not, thus raising the question of whether MPO, once inside an
endothelial cell, has any measurable function that could affect
cellular integrity. To test this, intracellular oxidant generation was
determined by monitoring the oxidation of nonfluorescent DHR to
fluorescent rhodamine. By UV light microscopy, HUVECs incubated with
MPO in the presence of DHR showed the generation of oxidants, displayed
as green cytoplasmic fluorescence (Figure 6B)
, whereas cells incubated with DHR
alone did not (Figure 6A)
. As analyzed by flow cytometry, MPO
incubation in the presence of DHR for 6 to 24 hours induced an increase
of fluorescence intensity in a dose- and time-dependent manner (Figure 6, E–G)
. MPO increased oxidant levels by 60% with 1 µg/ml, by 99%
with 5 µg/ml, and by 136% with 25 µg/ml treatment for 6 hours and
by 349% with 25 µg/ml treatment for 24 hours (Figure 6, F and G)
.
Using fluorometric methods, lysates of HUVECs treated with 25 µg/mg
MPO and DHR for 6 hours showed a 1.83-fold increase in oxidant
production. Similar to HUVECs, SAECs incubated with MPO in the presence
of DHR showed the generation of oxidants, displayed as green
cytoplasmic fluorescence by UV light microscopy (Figure 6D)
, whereas
cells incubated with DHR alone did not (Figure 6C)
. As measured by flow
cytometry, oxidant levels increased by 208% with 25 µg/ml MPO
treatment for 6 hours. Using fluorometric methods, lysates of SAECs
treated with 25 µg/ml MPO for 6 hours showed a 1.78-fold increase. If
MPO treatment results in increased levels of reactive oxygen species
leading to increased levels of
H2O2, then addition of
catalase would result in reduced oxidant levels. Catalase directly
catalyzes decomposition of
H2O2 to ground-state
O2.39
Although catalase is not cell
permeable, H2O2 can freely
diffuse across cell membranes and therefore would be available for
interaction with catalase in the tissue culture medium.40
Addition of catalase significantly blocked MPO-induced increases in
reactive oxygen species (Figure 6, F and G)
, in contrast to 1 to 10
µg/ml of PR3 for 6 to 24 hours, which did not induce generation of
intracellular oxidants in either HUVECs or SAECs (data not shown). The
data indicate that MPO is functionally active once internalized into
HUVECs or SAECs.
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Discussion |
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Internalization of inactive PR3 seems to be linked with activation of apoptosis. The mechanism of PR3 internalization may be unique, compared to that of MPO; however, a systematic study will be required to confirm these initial observations.
We have demonstrated that the C-terminal domain of the PR3 molecule
carries an apoptosis-inducing function, independent of its proteolytic
function. The data imply that inactive PR3 may exert its action through
penetration into the endothelial cell cytoplasm and/or nucleus and
perhaps even the nucleolus. It now becomes obvious that understanding
the mechanism of PR3-induced effects will be dependent on elucidation
of natural PR3-interacting proteins. Some known substrates of PR3 are
HSP28, nuclear factor-B, and Sp1, proteins found in both the
cytoplasm and the nucleus.21-23
If inactive PR3 were to
complex with these proteins, it is feasible that their normal functions
could be altered, even in the absence of proteolytic cleavage. In
support of this proposal some parallels can be drawn with the
extensively studied protease, granzyme B, a cytotoxic T-cell granule
protein with high homology to the neutrophil PR3. Characterization and
identification of substrates demonstrated that granzyme B recognizes an
80-kd nuclear protein and cytoplasmic proteins of 50, 94, and 69
kd.41
The 69-kd protein bound both the active and zymogen
forms of granzyme B. The C-terminal fragment of PR3 has an interesting
structure that may facilitate protein interactions and contribute to
its pathogenic effects. The only -helical domain in the PR3 molecule
is one at the C-terminal end of the molecule. The helix is downstream
of a hinge region, facilitating rotation and movement of the helix in
an arm-like manner, which would allow this domain to interact with
other proteins.
There is accumulating evidence that some neutrophil granule proteins have dual functions. Cathepsin G has a death function associated with the C-terminal portion of the protein, which has no proteolytic activity.42 Interestingly, the amino acid sequence of this region of cathepsin G is highly homologous to PR3-C, the region of PR3 containing the apoptosis-inducing function. Elastase has a death function separable from and independent of its proteolytic activity.43 It has recently been shown that azurocidin is internalized into endothelial cells and is targeted to mitochondrial compartments of the cell, but not in the nuclear compartment.44 Interestingly, internalized azurocidin markedly reduces growth factor deprivation-induced apoptosis, and the authors have suggested that uptake of exogenous azurocidin contributes to the sustained viability of endothelial cells in the context of locally activated neutrophils. The question arises of whether azurocidin would block PR3-induced apoptosis, thus revealing complexities of the balance/counterbalance mechanisms of the neutrophilic system never before realized.
Endothelial cells have been shown to internalize two other neutrophil granule proteins, lactoferrin45 and azurocidin.46 Lactoferrin is thought to enter the endothelial cell once complexed to binding sites on the cell surface for cationic proteins, whereas azurocidin binds to endothelial cell surface proteoglycans. However, neither of these two proteins are proteases. Proteases that are internalized by endothelial cells include renin and tissue plasminogen activator, both of which are internalized through binding to the mannose receptor.47,48 The mechanism of PR3 internalization has not been determined, however Taekema-Roelvink and co-workers49 have shown that PR3 interacts with a 111-kd membrane molecule of HUVECs. Further identification of PR3 binding molecules is currently under investigation.
MPO internalization has important implications in pinpointing mechanisms involved in oxidative damage in vivo. MPO uses H2O2 to produce diffusible cytotoxic oxidants.14 Internalization of MPO resulted in increased oxidant production through a catalase-inhibitable reaction. This means that MPO substrate H2O2 was present in these cells, in the absence of any other cytokine treatment. Vascular endothelial cells have been shown to produce basal levels of H2O2 at an intracellular site in the vicinity of peroxisomes and at a second site near the cell surface that is inaccessible to intracellular catalase.50 It has been shown also that cultured cells have as much as threefold greater rate of release of H2O2, in certain culture mediums.46 Therefore, it is highly possible that endothelial cells of vessels have low levels of intracellular H2O2. Titration of cellular H2O2 by MPO can result in decreased production of hydroxyl radical, thereby minimizing cell injury.51 H2O2 can react with superoxide anion to form the highly reactive hydroxyl radical. Purified MPO has been shown to strongly inhibit hydroxyl radical production in a concentration-dependent manner.52 Although MPO may have a protective effect in certain cases, MPO can react with H2O2 to form hypochlorous acid (HOCl), which is a more potent oxidant than H2O2, and excess hypochlorous acid formation can result in tissue damage. Active MPO has been shown to be a component of human atherosclerotic lesions,53 and internalization of MPO could potentate this disease process.
In summary, our data indicate the release of granule proteins by activated neutrophils and monocytes during acute inflammation may result in direct toxic effects by previously unrecognized mechanisms. The evidence indicates that PR3 enters endothelial cells and induces apoptosis through a function localized to the C-terminal death domain. We are very interested in understanding how C-PR3 is activating apoptosis. Exploration of the pathway(s) used by this fragment will require purified peptide. Efforts to generate this reagent have thus far been unsuccessful, because the C-PR3 fragment kills many of the expression systems that we have tried to use. In those systems that were not killed, we have yet to get purified protein. Studying the effects of internalization of neutrophil granule proteins or their fragments may well prove to be very important in understanding mechanism(s) of injury during acute inflammation.
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Supported by National Institute of Diabetes and Digestive and Kidney Diseases grant DK 40208.
Accepted for publication October 27, 2000.
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