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Release and Activation of Matrix Metalloproteinase-9 During In Vitro Mechanical Compression in Hypertrophic Scars
Filippo Renò, PhD;
Paola Grazianetti, PharmB;
Maurizio Stella, MD;
Gilberto Magliacani, MD;
Carla Pezzuto, MD;
Mario Cannas, MD
Arch Dermatol. 2002;138:475-478.
ABSTRACT
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Objective To investigate induction of matrix metalloproteinases (MMPs) during
mechanical compression of hypertrophic scars. Mechanical pressure blocks hypertrophy
inducted on extracellular matrix in scars by a mechanism that involves MMP-2
(gelatinase A) and MMP-9 (gelatinase B).
Design We assayed conditioned media obtained from normotrophic and hypertrophic
scars during 24 hours of in vitro mechanical compression using gelatin zymography.
Setting Scars from various areas of the bodies of hospitalized patients.
Patients We obtained 3 normotrophic and 7 hypertrophic biopsy specimens from
10 patients (5 men and 5 women).
Intervention In vitro compression at a pressure of 35 mm Hg/cm2 for 24
hours.
Main Outcome Measures Vitality of scars was analyzed by means of lactic dehydrogenase test;
medium samples were collected for zymographic analysis of MMP activity.
Results We found MMP-2 in basal (uncompressed) samples from normotrophic and
hypertrophic scars. Mechanical compression induced MMP-9 release and activation
(range, 86.7%-78.7%) in hypertrophic scars after 4 hours.
Conclusion Production, release, and activation of MMP-9 in hypertrophic scars could
be an effector mechanism responsible for hypertrophy regression induced by
mechanical compression.
INTRODUCTION
HYPERTROPHIC scars (HSs) result from alterations due to skin injuries,
especially burns, in the normal processes of cutaneous wound healing. Hypertrophic
scars are characterized by excessive deposition of fibroblast-derived extracellular
matrix (ECM) proteins, especially collagen, in the derma during a long period1 by means of persistent inflammation and fibrosis.2 The last phase of normal wound healing consists of
a systematic dissolution of granulation tissue and its replacement with a
more organized and elastic ECM.3 This process
seems to be altered in HSs.
The major effectors of ECM degradation and remodeling belong to a family
of structurally related enzymes called matrix metalloproteinases (MMPs).4 The MMPs are secreted in an
inactive proform requiring activation by means of proteolytic cleavage. Their
activity is zinc dependent and is inhibited by members of the family of tissue
inhibitors of metalloproteinases.5 The activity
of MMPs is regulated at multiple levels, and resident cells of a tissue typically
do not produce MMPs unless they are needed for remodeling.6
During cutaneous wound repair, the activity of MMP-2 (gelatinase A; 72-kd
type IV collagenase) and MMP-9 (gelatinase B, 92-kd type IV collagenase) persists
after wound closure and seems to be important in the remodeling process.7
Hypertrophy after burning is still treated mostly by means of elastocompression,
using elastic garments. The overall effect of pressure is an acceleration
of the remodeling process that, in the absence of pressure bandaging, can
occur several years after the formation of the HS. The mechanisms responsible
for hypertrophy remission after such a technique are still poorly understood,
although the interval of therapeutically useful pressure has been identified
as ranging from 10 to 35 mm Hg/m2.8
To our knowledge, no studies have investigated the role of MMPs in the regression
process induced in HS by elastocompression. In this in vitro study, we investigated
the presence of MMP-2 and MMP-9 in culture media conditioned with HSs and
normotrophic scars (NSs) and the effect of mechanical compression on MMP expression
and activation.
PATIENTS AND METHODS
We used 10 burn scars taken from different areas of the body. The biopsy
specimens (rectangular sections of 1 x 0.8 cm) were obtained after informed
consent from 10 patients (5 men and 5 women; mean [±SEM] age, 27 ±
13 years) undergoing surgery.
We assessed the clinical stage of each scar on the basis of macroscopic
observation before surgery and later by means of histological findings.
We examined NSs (n = 3) obtained from patients whose lesions underwent
optimal healing, and HSs (n = 7). Every sample was weighed and divided into
2 subspecimens, one to undergo compression and the other to serve as a control
sample. In vitro compression was performed using an electromechanical load
transducer (Instron 5564; Instron Corporation, Canton, Mass). The specimens
destined for compression were placed in an organ chamber containing 20 mL
of serum-free RPMI 1641 medium plus penicillin (100 U/mL) and streptomycin
(100 µg/mL), inserted into a thermostatic bath at 37°C, oxygenated,
and compressed using a metal punch. A pressure equal to 35 mm Hg was applied
to the section surface (skin side) for 24 hours. Control samples were maintained
in serum-free RPMI 1641 plus antibiotics at 37°C in an atmosphere of 5%
carbon dioxide.
Medium samples were taken from both subspecimens (1 mL) at the beginning
of the constant compression (pressure stabilized after 5-10 minutes at 35
mm Hg; time 0), and at 0.5, 1, 2, 4, and 24 hours. Before storing, we immediately
analyzed aliquots of all samples to check the vitality of the scar tissue
by using a commercial kit with a sensitivity of 3 to 5 U/L to evaluate the
amount of lactate dehydrogenase (LDH) (LDH-optimized UV method; Sigma-Aldrich
Corp, St Louis, Mo). We obtained the readings at 340 nm using a spectrophotometer
(Beckman DU-68; Beckman, Milan, Italy).
The samples were stored at -80°C and used for measurements
of MMP activity by means of zymography. Latent and active gelatinases were
detected by means of zymogram analysis using a combination of sodium dodecyl
sulfate and polyacrylamide gels copolymerized with 0.2% gelatin.7
These enzymes become dissociated from tissue inhibitors of metalloproteinases
by the presence of sodium dodecyl sulfate during electrophoresis. Removal
of sodium dodecyl sulfate after electrophoresis allows the proenzymes to renature
in an active or partially active conformation, which permits their detection
and the detection of lower-molecular-weight–activated forms. In brief,
conditioned media from control and compressed samples were mixed with sample
buffer and underwent electrophoresis directly without boiling or reduction.
After electrophoresis, sodium dodecyl sulfate was extracted from the polyacrylamide
gel using Triton X-100 (Sigma, Milan, Italy), and the gel was incubated in
a solution of 0.05M Tris (pH, 7.5) containing 5mM calcium chloride and 5mM
zinc chloride at 37°C overnight. Gels were stained with Coomassie brilliant
blue R-250 and then destained. Proenzyme and active gelatinase were detected
as clear bands against the blue background of the stained gelatin. Positive
controls for gelatinase A and B (Chemicon International, Inc, Temecula, Calif)
were used to distinguish the 2 enzymes from their activated forms. Unconditioned
medium samples were used as a negative control.
We performed a densitometric analysis of the bands seen on gels using
NIH Image 1.62 software (National Institutes of Health, Bethesda, Md). We
used SPSS software for Windows (SPSS Inc, Chicago, Ill) for the statistical
analysis of data. The t test was used for data; P<.05
indicated statistical significance.
RESULTS
We used gelatin zymography to monitor the activity of MMP-2 and MMP-9
in RPMI 1641 medium that was conditioned for 24 hours with biopsy specimens
obtained from burn scars. In the basal (uncompressed) condition, pro–MMP-2
was present at a barely detectable level in NS and at significantly higher
levels in HS (P<.001) (Figure 1 and Figure 2A).
The MMP-9 was undetectable in NS and HS in the basal condition, even if MMP-9
activity was present only in 1 HS sample (Figure 2A). The analysis of conditioned media obtained from compressed
specimens showed that pro–MMP-2 presence was almost undetectable in
NS and HS, with only 1 exception in an HS that already showed a strong MMP-2
activity in the uncompressed sample. In the same samples, the presence of
pro–MMP-9 and MMP-9 activities was observed mainly in HS (Figure 1 and Figure 2B).
Compression also induced the release of a small amount of pro–MMP-9
and MMP-9 in NS (Figure 1 and Figure 2B). The mean percentage of activation,
calculated as a percentage of the activated form divided by the total MMP-9
activity (proenzyme and activated form) for all 7 HSs examined, was 82.6%
(range, 78.7%-86.7%). The expression and activation of MMP-9 induced by the
mechanical compression in HS biopsy specimens was also time dependent (Figure 3). The active form of MMP-9 in compressed
specimens was detectable starting from 4 hours, and its presence increased
by 1200% after 24 hours. The MMP-9 mean percentage of activation after 4 hours
was 86.1% (range, 82.2%-88.4%), slightly but not significantly higher than
the 24-hour value. The mean (±SE) release of LDH measured after 24
hours in basal NS and HS samples (controls) was 35.9 ± 7.7 and 38.0
± 11.1 U/L, respectively. Compression caused a statistically insignificant
increase in LDH release (NS, 44.8 ± 17.2; HS, 51.1 ± 8.1), indicating
that no significant tissue damage occurred.
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Figure 1. Densitometric analysis of matrix
metalloproteinase (MMP)–2 and MMP-9 activity in normotrophic (NS; n
= 3) and hypertrophic scars (HS; n = 7) in the basal (uncompressed) condition
and after 24 hours of compression. Data are represented as mean ± SEM.
OD indicates optical density expressed in arbitrary units (AU). The MMP-2
activity value observed in 1 HS compressed sample was not indicated.
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Figure 2. Representative substrate gelatin
zymography of normotrophic (NS) and hypertrophic (HS) scar–conditioned
media. Media were conditioned with normotrophic (N1) and hypertrophic (HS1-4)
specimens for 24 hours in the basal condition (A) and in the compressed condition
(B). MMP indicates matrix metalloproteinase.
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Figure 3. Representative substrate gelatin
zymography of time-dependent release and activation of matrix metalloproteinase
(MMP)–9 in a hypertrophic (HS1) specimen during compression. Medium
samples have been collected at different time points (0-24 hours). The appearance
of MMP-9 activity is evident after 4 hours of compression. In this sample,
MMP-2 activity was also present.
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COMMENT
Pressure therapy is generally accepted as one of the best noninvasive
techniques for preventing and controlling hypertrophic scarring after burn
injury. This technique appears to arrest or suppress the production of additional
hypertrophic tissues and to enhance the natural remodeling process that occurs
long after the initial injury.9 Although this
method is efficient, its mechanisms of action are not known. Knowledge of
these mechanisms could result in better clinical treatment of burn injury.
Pressure therapy presents some adverse effects when used for extensive burns
or in children,10 and therefore the identification
of its effector mechanism(s) could lead to an alternative pharmacological
treatment of these patients. A previous study suggested that pressure accelerates
the remission phase of the postburn repair process by inducing ECM remodeling
and the disappearance of  -smooth muscle -actin expressing cells.11 Histological analysis of untreated HS showed concentric
nodular collagen fiber heaps in the deep dermis, whereas in HS treated by
elastocompression, the remodeling process was evident in the organization
of collagen, elastin, and fibrillin deposits.9, 11
Hypertrophy reduction has also been addressed to an ischemic process characterized
by cellular damage and collagen synthesis reduction,12
but this effect seems to be relevant only for prolonged treatment.9 In our in vitro model, this effect, if present, was
minimized by the oxygenation of the samples throughout the pressure treatment.
Moreover, the results of the LDH test showed no significant difference between
the viability of basal and compressed samples after 24 hours.
The major effector enzymes involved in the remodeling of the ECM are
MMPs.4 In particular, 2 of these enzymes, MMP-2
and MMP-9, seem to be involved in the remodeling phase of wound healing.7 In a previous study, MMP-2 activity was significantly
elevated in HS, whereas MMP-9 activity was undetectable13;
however, no studies have reported their possible involvement in the hypertrophy
remission induced by elastocompression. In our in vitro model, we observed
the presence of pro–MMP-2 in basal conditions in HS-conditioned medium,
whereas it was undetectable in samples obtained from NS. When HS samples were
compressed, MMP-9 activity appeared in a time-dependent fashion, whereas MMP-2
activity disappeared. At the same time in the NS samples, we found a light,
almost undetectable presence of MMP-2 and MMP-9 induced by compression. Matrix
metalloproteinase-9 can degrade native type IV and V collagens, elastin, fibronectin,
and denatured collagen of all types,14 and
its expression is induced by growth factors such as interleukin 1 and
transforming growth factor .15 The activation of MMP-9 is
believed to be induced by plasmin generated on the cell surface,16
but no information exists about this system in HS. In a previous study, we
observed that in vitro mechanical compression induced release of interleukin
1 and prostaglandin E2 in HS.17
Thus, in our model, interleukin 1 released by compression may induce
MMP-9 synthesis. The presence and activity of tissue inhibitors of metalloproteinases
could be checked in this model to understand whether they play a role during
the pressure-induced activation of MMP-9.
Given its broad activity against many substrates, the presence of MMP-9
could partially account for the ECM remodeling observed in pressure-treated
HS11 and for the remission of hypertrophy observed
in HS treated by elastocompression.8 However,
further investigation is necessary to validate this effect of compression
in an in vivo model and to identify the upstream events that trigger interleukin
1 release and MMP production, release, and activation.
AUTHOR INFORMATION
Accepted for publication July 9, 2001.
This study was supported by Fondazione Piemontese per gli Studi e le
Ricerche sulle Ustioni, Turin, Italy.
Corresponding author and reprints: Mario Cannas, MD, Human Anatomy
Laboratory, Medical Sciences Department, University of Eastern Piedmont "A.
Avogadro," Via Solaroli 17, 28100 Novara, Italy (e-mail: cannas{at}med.unipmn.it).
From the Human Anatomy Laboratory, Medical Sciences Department, University
of Eastern Piedmont "A. Avogadro," Novara (Drs Renò, Grazianetti, and
Cannas), and the Plastic Surgery Division and Burns Center, Orthopedic Traumatological
Center, Turin (Drs Stella, Magliacani, and Pezzuto), Italy.
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