Biol Reprod Track the topics, authors and articles important to you
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

BOR - Papers in Press, published online ahead of print November 14, 2007.
Biol Reprod 2007, 10.1095/biolreprod.107.063024
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
78/3/521    most recent
biolreprod.107.063024v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow My Folders
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.
Agricola
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.
BIOLOGY OF REPRODUCTION 78, 521–528 (2008)
DOI: 10.1095/biolreprod.107.063024
© 2008 by the Society for the Study of Reproduction, Inc.

Female Reproductive Tract; 

Regulation of Elastolytic Proteases in the Mouse Vagina During Pregnancy, Parturition, and Puerperium1

Cecilia K. Wieslander , Spyridon I. Marinis , Peter G. Drewes , Patrick W. Keller , Jesús F. Acevedo , and R. Ann Word 2

Division of Urogynecology and Reconstructive Surgery, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center, Dallas, Texas 75390

ABSTRACT

Recent evidence indicates that failure of elastic fiber assembly and synthesis is involved in the pathophysiology of pelvic organ prolapse in mice. It has been long been hypothesized that parturition-induced activation of proteases in the vaginal wall and its supportive tissues may contribute to pelvic organ prolapse in women. In this investigation, we determined the expression of matrix metalloproteases with elastase activity (matrix metalloproteinase [MMP] 2, MMP9, and MMP12) and their inhibitors in the vaginal wall of nonpregnant, pregnant, and postpartum mice. Data obtained using mRNA levels and enzyme activity measurements indicate that MMP2, MMP9, and 21- to 24-kDa caseinolytic serine proteases are regulated in vaginal tissues from pregnant and postpartum mice. Although suppressed during pregnancy and the early postpartum time period, MMP2 and MMP9 enzyme activities are increased after 48 h, a time when mRNA levels of protease inhibitors (tissue inhibitor of MMP2 [Timp2], cystatin C [Cst3], and alpha-1 antitrypsin [Serpina1]) are decreased. We conclude that recovery of the vaginal wall from pregnancy and parturition requires increased elastic fiber assembly and synthesis to counteract the marked increase in elastolytic activity of the postpartum vagina.

elastic fibers, matrix metalloprotease, pelvic organ prolapse, vagina, zymography

INTRODUCTION

Pelvic organ prolapse is a common condition that negatively impacts the quality of life of millions of women [1]. It has been recognized that the process of pregnancy, labor, and delivery is associated with the development of pelvic organ prolapse [2, 3]. However, the mechanisms by which pregnancy and parturition lead to failure of pelvic organ support are not known. Furthermore, the mechanisms that mediate the delayed manifestations of childbirth-associated injuries of the pelvic floor during childbirth are not understood. It has been hypothesized that parturition-induced injury of the vaginal fibroelastic tissue and activation of proteases in the vaginal wall and its supportive tissues contributes to pelvic organ prolapse in women [4–6]. The specific proteases involved in the vagina after parturition are not well defined, and there is little information regarding the regulation of extracellular matrix degradation in the vaginal wall postpartum.

Recently, two models of pelvic organ prolapse have been described, both of which involve defective synthesis (lysyl oxidase like-1, Loxl1) or assembly (fibulin-5, Fbln5) of elastic fibers. Loxl1 knockout mice appear to have normal pelvic organ support until after pregnancy and parturition, when animals exhibit prolapse of the pelvic organs [7]. Recent results indicate that a burst of elastic fiber assembly and cross-linking occurs in the vaginal wall postpartum, and that synthesis and assembly of elastic fibers are crucial for recovery of pelvic organ support after vaginal delivery [8]. Based on the relationship between failed elastic fiber synthesis and pelvic organ prolapse, we suggest that changes in elastic fiber degradation occur in the vaginal wall after parturition, thereby leading to failure of pelvic organ support in animals with the inability to synthesize new elastic fibers. Although pelvic floor connective tissues in women and mice are exposed to different degrees of mechanical forces, and mechanisms of pelvic organ prolapse may be unique in women, several investigations have demonstrated increased protease activity in vaginal wall connective tissue of women with prolapse [4, 9, 10]. Thus, the study of proteases that are known to degrade elastin may be relevant to the pathophysiology of prolapse in humans.

A number of different proteases have been implicated in the proteolytic degradation of elastic fibers and collagens, most prominent among which are members of the matrix metalloproteinase (MMP) family. MMPs form a subfamily of Ca2+-dependent zinc enzymes expressed predominantly in connective tissue and bone marrow cells [11, 12]. The enzyme family includes collagenases, gelatinases, and stromelysins, as well as membrane-type MMPs, matrilysin (MMP7), and metalloelastase (MMP12). MMPs can cleave virtually all protein components of the extracellular matrix (ECM). MMPs are produced as zymogens containing a secretory signal sequence and a propeptide of which proteolytic cleavage is required for MMP activation [13].

In this investigation, we tested the hypothesis that tissue remodeling of the vaginal wall after parturition involves activation of MMPs that exhibit elastolytic activity and determined the expression of matrix metalloproteases with elastase activity (MMP2, MMP9, and MMP12) and tissue inhibitors of elastases in the vaginal wall of nonpregnant, pregnant, and postpartum mice.

MATERIALS AND METHODS

Mice and Materials

Animals were housed under a 12L:12D (lights on, 0600–1800 h) cycle at 22°C. All mice used in the study were of C3BL/6J (The Jackson Laboratory, Bar Harbor, Maine) or mixed strain (C57BL/6 x 129SvEv). Timed-pregnant animals were obtained by housing the females with males for 4–6 h and checking midday for vaginal plugs. Plug day was considered Day 0. Birth occurred in the early morning hours (0200–0600 h) of Day 19. All studies were conducted in accordance with the Guide for the Care and Use of Laboratory Animals using protocols approved by the Institutional Animal Care and Use Committee. Unless indicated otherwise, all reagents were purchased from Sigma Aldrich (St. Louis, MO).

Tissue Processing

Mice were killed at nonpregnant, pregnant, parturient, or postpartum time points. After disarticulation of the pubic symphysis, the uterine horns, bladder, cervix, and vagina were dissected down to the perineal skin. Using microinstruments and a dissection microscope, the uterine horns were removed at the level of the cervicovaginal junction. Perineal skin was removed, and the bladder and urethra were dissected from the anterior vaginal wall. The wet weight of vagina and cervix was determined. Thereafter, the cervix was removed from the vaginal tube and weighed. For RNA analysis, tissues were stored at –20°C in RNALater (Ambion, Austin, TX). Tissues were frozen in liquid N2 for analysis of enzyme activity.

Real-Time PCR

Tissues were minced and homogenized in 4 M guanidinium isothiocyanate buffer, layered over 5.7 M cesium chloride, and centrifuged overnight at 237 000 x g to extract RNA. Concentration of RNA was measured and purity confirmed by spectroscopy. Reverse transcription reactions were conducted with 2 µg total RNA in a reaction volume of 20 µl. Each reaction contained 10 mM dithiothreotal (DTT), 0.5 mM deoxynucleotide triphosphates (dNTPs), 0.015 µg/µl random primers, 40 U RNase inhibitor (Invitrogen #10777–019; Invitrogen, Carlsbad, CA), and 200 U reverse transcriptase (Invitrogen #18064–014; Invitrogen). Primer sequences for amplifications were chosen using published cDNA sequences and the Primer Express program (Applied Biosystems, Foster City, CA). Primers were chosen such that, when possible, the resulting amplicons would cross an exon junction, thereby eliminating false positive signals from genomic DNA contamination (Table 1). SYBR Green was used for amplicon detection. Gene expression was normalized to expression of the housekeeping gene beta-2 microglobulin (B2m). Positive controls (heart, lung, and spleen) were run on each plate as appropriate, and all assays included no template controls. All primer sets were tested to ensure that efficiency of amplification over a wide range of template concentrations was equivalent to that of B2m. Positive and negative tissue controls for each primer set were included in each reaction. PCR reactions were carried out in the ABI Prism 7000 sequence-detection system (Applied Biosystems). The reverse transcription product from 50 ng RNA was used as template, and reaction volumes (30 µl) contained 1 Master Mix (Applied Biosystems #4309155). Primer concentrations were 900 nM. Cycling conditions were: 2 min at 50°C, followed by 10 min at 95°C; then 40 cycles of 15 sec at 95°C, and 1 min at 60°C. A preprogrammed dissociation protocol was used after amplification to ensure that all samples exhibited a single amplicon. Levels of mRNA were determined using the ddCt method (Applied Biosystems) and expressed relative to an external calibrator present on each plate.


View this table:
[in this window]
[in a new window]
[Download PPT slide]
  		TABLE 1. Sequences of primers utilized for quantification of gene expression by real-time PCR.

MMP2 Protease Activity

Tissues used for protease activity determinations and zymography were dissected as described above. The vaginal tube was incised longitudinally, and the vaginal epithelium was removed by scraping with a scalpel. Thereafter, tissues were snap-frozen in liquid N2. To measure enzyme activity, tissues were thawed on ice, minced, and washed in PBS with 2 mM N-methylamine (an inhibitor of antielastase activity) until the supernatant was clear. Tissues were then homogenized in MMP2 assay buffer (AnaSpec EnzoLyte 520 MMP2 Assay Kit; AnaSpec, San Jose, CA) containing 0.1% Triton-X 100 (95x volume:tissue wet weight). Thereafter, the homogenates were centrifuged at 10 000 x g for 15 min at 4°C. The supernatant was used for determination of protease activity. Protein concentrations were determined using a BCA protein assay and standard curves of BSA in appropriate buffers. Pro-MMP2 enzyme was activated by incubating the tissue homogenates with 1 mM 4-aminophenylmercuric acetate (APMA) in MMP2 assay buffer for 15 min at 37°C. Enzyme activity was determined using a fluorescently labeled peptide (FRET peptide) as substrate and a standard curve of purified MMP2 as recommended by the manufacturer (AnaSpec EnzoLyte 520 MMP2 Assay Kit). Fluorescence intensity was measured at Ex/Em = 490 nm/520 nm every 5–10 min for 40–60 min. Selective protease activity was inhibited by incubating the samples for 30 min with 12.5 mM EDTA (MMP inhibitor), 12.5 nM tissue inhibitor of MMP2 (TIMP2), 25 mM iodoacetamide (IAMD, cysteine protease inhibitor), and 2.5 mM PMSF (serine protease inhibitor). EDTA and iodoacetamide were added to the reaction from 250 mM stock solutions in assay buffer. TIMP2 was added from a stock solution of 1 µM in assay buffer. PMSF was prepared as a 100-mM stock solution in ethanol. Thereafter, it was diluted to 50 mM in buffer and to a final concentration of 2.5 mM in the reaction. All control and inhibitor reactions received identical vehicle concentrations.

Gelatin Zymography

Samples (5 µg per lane) were applied to gelatin polyacrylamide minigels (10%; Invitrogen) in standard SDS loading buffer containing 0.1% SDS with no β-mercaptoethanol, and the samples were not boiled before loading. The gels were run at room temperature at 125 V. After electrophoresis, the gels were soaked in renaturing buffer (2.7% [v/v] Triton X-100 in distilled water) in a shaker for 30 min, with one change after 30 min to remove SDS. Next, the gels were soaked in developing buffer (50 mM Tris, 200 mM NaCl, 10 mM CaCl2, 0.05% Brij 35; pH 7.5) overnight at 37°C and stained with Coomassie brilliant blue-R 250 in 50% methanol and 10% acetic acid, followed by washing with distilled water for 1 min. Clear zones of lysis against a dark background indicated enzyme activity. Areas of lysis were quantified using a Fuji LAS 3000 image analysis system.

Casein Zymography

Tissue extracts were mixed with SDS-PAGE sample buffer (10% glycerol, 2% SDS, and 63 mM Tris; pH 7.0) without reducing agent and applied to nonreducing 4%–16% acrylamide gels containing 0.1% casein (NuPAGE; Novex, Encinitas, CA). Electrophoresis was carried out for 90 min at 125 V at room temperature, and resolved proteins were renatured in situ by immersing the gels in 2.7% (wt/v) Triton X-100 for 30 min at room temperature. The gels were then rinsed in zymogram developing buffer (Novex) for 30 min and incubated overnight at 37°C in the same buffer. Caseinolytic activity was visualized as negative staining against the blue background. Zymograms shown in this study are representative replicates selected from at least two experiments.

Statistical Analysis

Protease activity and mRNA levels were compared using SigmaStat software (Jandel Scientific, San Rafael, CA). A t-test, paired t-test, ANOVA, and Kruskal-Wallis ANOVA on ranks were used to compare groups, using nulliparous nonpregnant mouse vagina or early pregnancy mouse vagina as controls. P ≤ 0.05 was considered statistically significant.

RESULTS

Expression of MMPs with Elastase Activity in the Vagina During Pregnancy and Parturition

Expression of Mmp2, Mmp9, and Mmp12 mRNA was determined by real-time PCR in vaginal tissues from nonpregnant and pregnant mice during pregnancy, parturition, and the postpartum time period (Fig. 1). Mmp2 and Mmp9 transcripts were highly expressed in the vaginal wall, with levels of expression greater than that in mouse liver (Mmp2) or spleen (Mmp9). However, Mmp12 was not expressed in the vaginal wall (CT = 30) and was not regulated during pregnancy. Mmp2 mRNA levels were suppressed in the vaginal wall from animals in labor and immediately postpartum (2–4 h; Fig. 1A). Mmp2 mRNA was increased significantly in vaginal tissues from animals 12–24 h after delivery. Mmp2 mRNA levels returned to baseline by 48 h and were maintained for 2 wk postpartum. The gestational pattern of MMP9 expression was similar to that of Mmp2 in that Mmp9 mRNA levels were decreased significantly in the early postpartum time period (Fig. 1B). Unlike Mmp2, however, levels of Mmp9 mRNA remained suppressed for 12 h postpartum, increasing >10-fold by 24–48 h (Fig. 1B). These levels returned to nonpregnant levels within 1 wk after delivery.


Figure 1
View larger version (14K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 1. Regulation of Mmp2 and Mmp9 mRNA in the vaginal wall during pregnancy, parturition, and the postpartum time period. The relative abundance of Mmp2 (A), Mmp9 (B), and Mmp12 (C) mRNA was determined by real-time PCR in vaginal tissues from nonpregnant (NP, open bar), and pregnant mice at various time points in gestation (hatched bars) or postpartum (solid bars). Data is expressed relative to B2m and normalized to a nonpregnant vagina standard. Each data point represents mean ± SEM of 3–11 tissues at each time point. Early Pregnant, Days 8–12; In Labor, in labor after delivery of the first pup; *P ≤ 0.05 compared with Day 18 pregnant time point.

Regulation of MMP2 Enzyme Activity During Pregnancy, Parturition, and the Postpartum Time Period

Regulation of posttranslational processing of inactive proenzymes to mature active proteases is important in orchestrating extracellular matrix remodeling by metalloproteases [13]. To examine the effect of pregnancy on MMP2 enzyme processing, tissue extracts were obtained from nonpregnant, pregnant, and postpartum mice at various time points, and a fluorescently labeled substrate specific for MMP2 was used to determine enzyme activity. The substrate, however, may also be cleaved by MMPs 8, 9, 12, 13, and 14. APMA cleaves MMPs, thereby converting inactive proforms to active enzymes. Enzyme activity in the absence of APMA was utilized to assess the level of endogenous mature active enzyme within the tissue homogenate (Fig. 2A), and the activity of both pro- and active enzymes was determined by incubating tissue homogenates with APMA prior to the assay (Fig. 2B). Standard curves were also determined in the presence and absence of APMA. Endogenous MMP2 activity was significantly decreased in pregnant mice in labor and in the early postpartum time period (2–12h; Fig. 2A). MMP2 activity, however, increased significantly 48 h after delivery, returning to nonpregnant baseline levels by 4 wk. Interestingly, treatment with APMA resulted in decreased enzyme activity in vaginal tissue homogenates from nonpregnant, pregnant, and early postpartum mice (Fig. 2B). APMA, however, did not alter enzyme activity measurements of purified MMP2, suggesting that factors within the tissue homogenate inhibited APMA-activated MMP2 activity. In contrast, APMA resulted in either similar or increased enzyme activity in tissue homogenates from postpartum mice after 24 h. The results indicate that during pregnancy and immediately after delivery, endogenous MMP2 is suppressed, and that both active and proenzymes are increased in vaginal tissues from late postpartum mice (48 h).


Figure 2
View larger version (8K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 2. Regulation of MMP2 enzyme activity in vaginal tissues during pregnancy, parturition, and postpartum time period. A) Endogenous MMP2 enzyme activity was quantified in the absence of APMA in vaginal tissues from nonpregnant (NP, open bar) and pregnant mice at various time points in gestation (hatched bars) or postpartum (solid bars). In Labor, in labor after delivery of the first pup; *P ≤ 0.05 compared with nonpregnant. B) Total MMP2 activity was quantified in vaginal tissue homogenates from nonpregnant (NP, open bar) and pregnant mice at various time points in gestation (hatched bars) or postpartum (solid bars) after incubation with APMA (1 mM) for 15 min prior to quantification of MMP2 enzyme activity. Each data point represents mean +/– SEM of four to eight tissues at each time point. *P ≤ 0.05 compared with nonpregnant, **P ≤ 0.05 compared with endogenous activity at the corresponding time point.

MMP2 enzyme activity was characterized further in vaginal tissue extracts from nonpregnant and postpartum mice (Fig. 3). Because Zn2+ and Ca2+ are required for MMP2 activity, enzyme activity was determined in tissue extracts in the presence or absence of EDTA. As expected, EDTA resulted in marked inhibition of enzyme activity in tissue extracts from both nonpregnant and postpartum mice. Because MMP2 activity is regulated by TIMP2, a protein secreted in the extracellular matrix that binds to and inhibits MMP activity, tissue extracts from nonpregnant and postpartum mice were incubated with purified TIMP2, and enzyme activity was determined (Fig. 3). Although TIMP2 resulted in significant decreases in MMP2 activity in both tissue extracts, inhibition was more pronounced in vaginal tissues from postpartum animals (P ≤ 0.05), suggesting that more TIMP2 binding sites are available in the postpartum vagina.


Figure 3
View larger version (10K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 3. Effect of EDTA and TIMP2 on MMP2 activity in the vaginal wall. Tissue homogenates from nonpregnant (A) or postpartum (24 h, B) mice were incubated with vehicle (CTL), EDTA (12.5 mM), or TIMP2 (12.5 nM) for 30 min prior to quantification of MMP2 enzyme activity. Error bars represent SEM of four samples in each group. *P ≤ 0.05 compared with control; **P ≤ 0.001 compared with nonpregnant vaginal tissues.

MMP2 and MMP9 Enzyme Activity During Pregnancy, Parturition, and the Postpartum Time Period

Because MMP2 enzyme activity assays have significant cross-reactivity with other MMPs, gelatin zymography was used to confirm the MMP2 assay results and assess both pro- and active forms of MMP9 and MMP2 in vaginal tissues from nonpregnant and postpartum mice (Fig. 4). In vaginal tissues from nonpregnant mice, MMP9 activity was low, but detectable (using 5 ug of extract protein per lane) as a gelatinolytic protein of ~92 kDa (proMMP9). ProMMP9 was converted to active MMP9 (~86 kDa) by treatment with APMA. In nonpregnant mice, MMP2 was more abundant than MMP9 and was expressed as pro- (72 kDa), intermediate (68 kDa), and active (62 kDa) forms of the enzyme (Fig. 4). Interestingly, treatment of extracts from nonpregnant animals with APMA resulted in decreased amounts of proMMP2 and concomitant increased amounts of the intermediate form, but no increases in the mature 62-kDa form of MMP2.


Figure 4
View larger version (43K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 4. MMP9 and MMP2 enzyme activity in vaginal tissues from nonpregnant and postpartum mice. A) Gelatin zymography (5 µg per lane) was conducted in homogenates of vaginal tissues from nonpregnant or puerperial (48h postpartum) mice treated with (+) or without (–) APMA. APMA resulted in processing of proMMP9 (92 kDa) to active MMP9 (86 kDa) and proMMP2 (72 kDa) to 65- and 62-kDa enzymes. MMP9 is increased dramatically in vaginal tissues from postpartum mice compared with nonpregnant controls. B) Quantification of pro- (open bars) and active (closed bars) MMP9 and MMP2 in vaginal tissues from nonpregnant (NP) and postpartum (PP) mice. Error bars represent SEM of three samples in one experiment. The experiment was repeated with identical results. *P ≤ 0.05 compared with NP.

Both pro- and active forms of MMP9 and MMP2 were increased dramatically in the postpartum vagina (Fig. 4). In contrast with nonpregnant animals, MMP9 was the predominant protease in vaginal tissues from postpartum mice. In the presence of APMA, proMMP9 was converted to its active form, and proMMP2 was converted predominantly to the 62-kDa form. Serum is enriched in elastase inhibitors such as {alpha}-2-macroglobulin. To diminish the effect of these inhibitors on proteases in tissue homogenates, tissues were collected and rinsed in N-methylamine. To ensure that upregulation of MMP activity in the postpartum vagina was not due to reduced activity of elastase inhibitors, tissues from nonpregnant and postpartum (48 h) mice were analyzed in the absence of N-methylamine (Fig. 5). The magnitude of change in protease activity in the postpartum vagina was similar to that observed in Fig. 4. Thus, increased MMP2 and MMP9 in the postpartum vagina is not due to experimental effects of N-methylamine.


Figure 5
View larger version (28K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 5. Gelatinase activity in vaginal tissues from postpartum animals. Gelatin zymography was conducted in homogenates of vaginal tissues from nonpregnant and puerperal mice (48 h PP). Tissue homogenates were prepared without the elastase inhibitor N-methylamine. Data given in kDa.

Casein Zymography: Protease Activity in Vaginal Tissues from Nonpregnant and Postpartum Mice

To further characterize proteases of different substrate specificities, extracts of vaginal tissues from nonpregnant and postpartum animals were subjected to SDS-PAGE under non-reducing conditions in gels containing casein (Fig. 6). Caseinolytic activity was detected as clear bands. Caseinolytic proteases migrating at 98, 24, and 21 kDa were not observed in vaginal tissues from nonpregnant animals, but were markedly upregulated in tissues from postpartum mice. A caseinolytic protease migrating ~60 kDa was faint and not observed consistently on all casein zymograms. The identity of the 98-kDa protease was not MMP9, because its activity was not decreased by EDTA. The 24/21-kDa protease activity was not inhibited by EDTA or iodoacetamide (Fig. 6B). Significant inhibition (>70%), however, was observed with PMSF, suggesting that this protease was not a metalloprotease but rather a member of the serine protease family.


Figure 6
View larger version (27K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 6. Casein zymography of vaginal tissue extracts. A) Vaginal extracts (20 µg per lane) from nonpregnant (lanes 1–5) and postpartum (48 h, lanes 7–12) mice were applied to an SDS polyacrylamide gel that contained casein-blue (0.1%). After electrophoresis, the gel was incubated at 37° for 18 h. Molecular weight markers are indicated on the right. B) Zones of lysis in the 21- to 24-kDa range were quantified using densitometry of vaginal homogenates. Homogenates from postpartum mice were treated with IAMD (25 mM), EDTA (12.5 mM), or PMSF (2.5 mM) and compared with that of vehicle (Ctl). Error bars represent SEM of three samples. *P ≤ 0.05 compared with control.

Expression of Elastase Inhibitors in the Vagina During Pregnancy and Parturition

MMP2 and MMP9 activity is controlled not only by transcription, translation, and processing, but also by endogenous protein inhibitors within the tissue that bind to, and inhibit, protease activity. Tissue inhibitor of MMP1 (TIMP1) and TIMP2 are proteins that primarily regulate MMP9 and MMP2 enzyme activity, respectively. In the vaginal wall, Timp1 was less abundant than Timp2 (CT values of 26–27 compared with 20–21 in vaginal tissues from nonpregnant mice). Although MMP9 activity was increased in vaginal tissues of postpartum mice, Timp1 was also increased significantly (Fig. 7A). In contrast, expression of Timp2 was decreased 4-fold in vaginal tissues from animals 48 h postpartum (Fig. 7B), suggesting that increased MMP2 enzyme activity in the postpartum vagina may be, at least partially, due to a decrease in its inhibitor.


Figure 7
View larger version (10K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 7. Expression of Timp1 and Timp2 mRNA in vaginal tissues during pregnancy, parturition, and the puerperium. Real-time PCR was used to determine the relative abundance of Timp1 (A) and Timp2 (B) in vaginal tissues from nonpregnant (NP), pregnant (D18), and postpartum mice 24 (PP 24 h) and 48 h (PP 48 h) after delivery. Data represent mean ± SEM of five to seven tissues at each time point. *P ≤ 0.01 compared with nonpregnant, D18, or 24 h; **P ≤ 0.01 compared with nonpregnant.

Three protease inhibitors known to inhibit other elastases are cystatin C (CST3), {alpha}1-antitrypsin (SERPINA1), and secretory leukocyte protease inhibitor (SLPI) [14–18]. These inhibitors were highly expressed in the vaginal wall (CT = ~19 using 50 ng RNA). Cst3 mRNA levels increased progressively from early in pregnancy until Day 18, were maintained in labor and 24 h postpartum, and then returned to baseline levels 48 h after delivery (Fig. 8A). The pattern of expression of Serpina1 was similar to Cst3 except Serpina1 mRNA levels peaked on Day 18 of pregnancy and declined thereafter (Fig. 8B). Like Cst3, expression of Serpina1 was decreased significantly 48 h after delivery. Expression of Slpi was variable in the vaginal wall throughout pregnancy and the postpartum time period, suggesting that transcripts for this protease inhibitor may not be regulated during parturition (Fig. 8C).


Figure 8
View larger version (7K):
[in this window]
[in a new window]
[Download PPT slide]
  		FIG. 8. Expression of anti-elastase mRNA in vaginal tissues during pregnancy, parturition, and the puerperium. Real time PCR was used to determine the relative abundance of cystatin C (Cst3, A), Serpina1 (B), and secretory leukocyte protein inhibitor (Slpi, C) mRNA in vaginal tissues from pregnant and postpartum mice at various time points. Data is expressed relative to B2m. Each data point represents mean ± SEM of 4–11 tissues at each time point. 8–12, 16, and 18 indicate D; IL, in labor after delivery of the first pup; 24h, 24 h postpartum; 48h, 48 h postpartum; 1 wk, 1 wk postpartum; 2 wk, 2 wk postpartum. *P ≤ 0.05 compared with early pregnancy time point; {delta}P = 0.04, although not statistically significant using pairwise multiple comparisons.

Taken together, data obtained using mRNA levels and enzyme activity measurements indicate that MMP2, MMP9, and 21- to 24-kDa caseinolytic serine proteases are regulated in vaginal tissues from pregnant and postpartum mice. Although suppressed during pregnancy and the early postpartum time period, MMP2 and MMP9 activities are increased after 48 h, a time when mRNA levels of protease inhibitors (Timp2, Cst3, and Serpina1) are decreased.

DISCUSSION

Perhaps the most dramatic physiologic tissue remodeling process in all of biology occurs in the female reproductive tract during pregnancy, parturition, and the postpartum time period. Tissue remodeling requires two basic processes that change tissue architecture through alterations in the composition of the ECM. On one hand, remodeling requires synthesis and deposition of ECM proteins, whereas, on the other, proteolytic degradation of the ECM is also required. The vaginal wall undergoes continuous remodeling in response to varying levels of hormones in the estrus cycle [19–23] and during pregnancy [23]. Previously, we reported that elastic fiber synthesis and assembly is suppressed in the vagina during pregnancy, but rebounds significantly by 48 h after delivery [8]. In this investigation, we demonstrated that this upregulation of elastic fiber synthesis and assembly in the postpartum time period is accompanied by dramatic increases in proteases that target elastic fibers.

Regulation of MMP2 and MMP9 in the Vaginal Wall During Pregnancy and the Puerperium

Elastin-degrading proteases consist of cysteine proteases (cathepsins B, H, L, and K), serine proteases (e.g., neutrophil elastase), and a subset of MMPs (MMP2, MMP9, and MMP12). Because MMPs are believed to play central roles during both physiological and pathological matrix remodeling, in this investigation we focused on the regulation of MMP2, MMP9, and MMP-12 and elastase inhibitors. We found that MMP2 and MMP9 levels were suppressed and antielastase mRNA was increased in the vagina during late pregnancy and the early postpartum time period. These findings suggest that adaptations of the vaginal wall during pregnancy include protection of elastic fibers from degradation. Mmp2 and Mmp9 mRNA and enzyme activity increased, however, in vaginal tissues from postpartum mice at later time points (12–48 h), a time in which antielastase and Timp2 mRNA levels were decreased. These results suggest that the postpartum vagina may be particularly vulnerable to elastic fiber degradation. It should be noted, however, that simultaneous increases in elastic fiber synthesis and assembly 24–48 h after delivery counteract the degradative pathway, resulting in a net increase in elastic fibers after parturition [8]. The significant increase in desmosine content (an index of mature cross-linked elastin) in vaginal tissues of parous mice compared with virginal animals [8] is consistent with this hypothesis.

Activation of MMP2 in Vaginal Tissues

In the vaginal wall, both pro- and active forms of MMP2 and MMP9 were increased significantly postpartum. Increased MMP2 activity appeared to be due to increased MMP2 synthesis and decreased expression of Timp2. In contrast, increases in MMP9 activity appeared to be predominantly due to increased synthesis of the enzyme because Timp1 levels were also increased in the postpartum vagina. Increased TIMP1 may serve to limit MMP9 activity in the postpartum time period.

Most mammalian cells produce proMMP2 tightly complexed with TIMP2 [24, 25]. The stoichiometry of the components of this complex, however, is differentially regulated in reproductive tissues during pregnancy [19]. In most resting tissues, proMMP2 exists in an equimolar ratio of TIMP2, and proMMP2*TIMP2 complexes show autolysis upon activation with APMA [26, 27]. It was surprising to find that, in vaginal tissues from nonpregnant and pregnant mice (but not postpartum mice), APMA-activated MMP2 activity was significantly less than endogenous MMP2 activity. There are several possible explanations for this phenomenon. Previous studies indicate that although the presence of TIMP2 in the complex does not prevent chemical activation of the enzyme by APMA [28], binding of APMA-activated MMP2 to free TIMP2 prevents the rapid autolytic conversion induced by APMA [29] and thereby inhibits APMA-activated enzyme activity [28]. In addition, recent studies using purified free enzymes and MMP2*TIMP2 complexes indicate that the free enzyme can be activated rapidly and fully by APMA, but that identical treatment of MMP2*TIMP2 complexes with APMA results in significantly slower and partial activation [29]. Results in this investigation indicating that expression of TIMP-2 is increased significantly in vaginal tissues from nonpregnant or pregnant mice and that resting MMP2 activity in these tissues is less sensitive to inhibition by TIMP-2 suggest that native TIMP2 is complexed with proMMP2 in vaginal tissues from nonpregnant and pregnant mice, and that when this enzyme complex is activated, TIMP2 is free to interact with the converted MMP2, substantially reducing the specific activity of the generated enzyme [29]. Overall, the results support the idea that different activities of MMP2 in vaginal tissues from nonpregnant/pregnant and postpartum mice are due to differences in MMP2 enzyme and its inhibitor.

Increased Expression of Other Proteases in the Postpartum Vagina

Casein zymography revealed proteases of 98, 62–65, and 21–24 kDa. The serine protease inhibitor PMSF inhibited the 21- to 24-kDa enzymes. Although the identity of the 21- to 24-kDa proteases is not readily apparent, several possibilities exist. A cathepsin G-like serine protease of similar molecular weight has been reported in injured human and rat corneas [30] and a caseinolytic protease of 21- to 24-kDa that degrades IGFBP5 has been reported in human fibroblasts [31]. To our knowledge, this is the first report of a regulated serine protease in the vagina, and work is ongoing to identify the specific protein. Upregulation of both MMPs and serine proteases of different substrate specificities in the postpartum vagina further emphasizes the complexities of the remodeling process as the vaginal wall recovers from pregnancy and parturition.

Elastase Inhibitors in the Vaginal Wall

Elastase activity is controlled on at least three levels: transcription, proteolytic activation of zymogen forms (proMMPs), and inhibition of active enzymes by a host of natural inhibitors. Several inhibitors are expressed in vaginal epithelial cells and secreted into the underlying supportive ECM [32–35]. For example, CST3 is a secreted inhibitor of cysteine proteinases [16] and has been shown to regulate cysteine protease activity during bone resorption and other tissue remodeling processes [36]. Lack of CST3 and increased cysteine proteinase activity is associated with numerous pathological conditions associated with ECM degradation, including atherosclerosis [16] and invasive cancer [32, 37]. Although CST3 is expressed in virtually every tissue, mRNA levels vary greatly between different tissues under various conditions [38]. For example, estrogen increases CST3 expression in the vaginal wall [38]. Neutrophil elastases and other elastases are controlled predominantly by endogenous inhibitors such as SERPINA1 and {alpha}-2-macroglobulin [39, 40]. Cst3 and Serpina1 mRNA levels increased in the vaginal wall, but expression of both antielastases declined to resting levels 48 h after delivery. The results suggest that increased expression of these protease inhibitors may protect the vaginal wall from substantial degradation during pregnancy, but that this protection is abrogated 48 h postpartum.

In summary, most MMPs are either not expressed or expressed at low levels in resting-state adult tissues [13]. However, numerous cytokines, growth factors, hormones, and physical cellular interactions provide stimuli that can rapidly induce MMP expression. MMP2 mRNA and enzyme activity were relatively high in vaginal wall tissues from nonpregnant mice, possibly because of the continuous remodeling processes that accompany dramatic changes in the size and morphology of this organ during the estrus cycle. We found that MMP activity is regulated in the vaginal wall during pregnancy, parturition, and the puerperium. During pregnancy and immediately postpartum, expression of MMP2 and MMP9 is suppressed while that of elastase inhibitors is increased in the vagina. Although MMP2 and MMP9 are increased in the vagina 48 h after delivery, the burst of fibulin-5, tropoelastin, and lysyl oxidase may counteract elastin degradation, resulting in net increases in elastic fiber deposition and recovery of the vagina from parturition [8]. Increased protease activity, together with deficient repair mechanisms involving regeneration of elastic fibers, may lead to overall degradation of the ECM in connective tissues of the pelvic floor and prolapse of the pelvic organs. Disruption of the balance between matrix degradation and repair in vaginal connective tissues after vaginal delivery may also lead to failure of pelvic organ support in women.

FOOTNOTES

1Supported by NIH AG 028048. Back

Correspondence: 2R. Ann Word, Division of Urogynecology and Reconstructive Surgery, Department of Obstetrics and Gynecology, University of Texas Southwestern Medical Center at Dallas, 5323 Harry Hines Blvd., Dallas, TX 75390-9032. FAX: 214 648 9242; e-mail: ruth.word{at}utsouthwestern.edu

Received: 25 May 2007.

First decision: 12 July 2007.

Accepted: 5 November 2007.

REFERENCES

  1. ; : Olsen AL, Smith VJ, Bergstrom JO, Colling JC, Clark AL. Epidemiology of surgically managed pelvic organ prolapse and urinary incontinence. Obstetrics and Gynecology 1997 89501–506[CrossRef][Medline]
  2. The Genealogy of Gynecology . . Ricci J. 1950Philadelphia: Blakiston Co;
  3. ; : Emge L and Durfee R. Pelvic organ prolapse: four thousand years of treatment. Clin Obstet Gynecol 1966 9997–1032[CrossRef][Medline]
  4. ; : Alperin M and Moalli PA. Remodeling of vaginal connective tissue in patients with prolapse. Curr Opin Obstet Gynecol 2006 18544–550[Medline]
  5. ; : Moalli PA, Shand SH, Zyczynski HM, Gordy SC, Meyn LA. Remodeling of vaginal connective tissue in patients with prolapse. Obstet Gynecol 2005 106953–963[Medline]
  6. ; : Moalli PA, Howden NS, Lowder JL, Navarro J, Debes KM, Abramowitch SD, Woo SL. A rat model to study the structural properties of the vagina and its supportive tissues. Am J Obstet Gynecol 2005 19280–88[CrossRef][Medline]
  7. ; : Liu X, Zhao Y, Gao J, Pawlyk B, Starcher B, Spencer JA, Yanagisawa H, Zuo J, Li T. Elastic fiber homeostasis requires lysyl oxidase-like 1 protein. Nat Genet 2004 36178–182[CrossRef][Medline]
  8. ; Drewes PG, Marinis SI, Acevedo J, Keller P, Schaffer JI, Word RA. Regulation of lysyl oxidase in the vaginal wall: Role of estrogen, progesterone, and pregnancy. From the 2005 26th Annual Scientific Meeting of The American Urogynecologic Society. J Pelv Med Surg 2005 11(suppl 1):S8–9
  9. ; : Chen B, Wen Y, Polan M. Elastolytic activity in women with stress urinary incontinence and pelvic organ prolapse. Neurourol Urodyn 2004 23119–126[CrossRef][Medline]
  10. ; : Phillips CH, Anthony F, Benyon C, Monga AK. Collagen metabolism in the uterosacral ligaments and vaginal skin of women with uterine prolapse. BJOG 2006 11339–46[CrossRef][Medline]
  11. ; : Rawlings ND and Barrett AJ. Evolutionary families of metallopeptidases. Methods Enzymol 1995 248183–228[Medline]
  12. ; Shapiro SD. A concise yet informative stroll through matrix metalloproteinases and TIMPs. J Cell Sci 2000 113(pt 19):3355–3356[Abstract]
  13. ; : Stamenkovic I. Extracellular matrix remodelling: the role of matrix metalloproteinases. J Pathol 2003 200448–464[CrossRef][Medline]
  14. ; : Overall CM and Dean RA. Degradomics: systems biology of the protease web. Pleiotropic roles of MMPs in cancer. Cancer Metastasis Rev 2006 2569–75[CrossRef][Medline]
  15. ; : Zeeuwen PL. Epidermal differentiation: the role of proteases and their inhibitors. Eur J Cell Biol 2004 83761–773[CrossRef][Medline]
  16. ; : Liu J, Sukhova GK, Sun JS, Xu WH, Libby P, Shi GP. Lysosomal cysteine proteases in atherosclerosis. Arterioscler Thromb Vasc Biol 2004 241359–1366[Abstract/Free Full Text]
  17. ; : Barrett AJ. The cystatins: a diverse superfamily of cysteine peptidase inhibitors. Biomed Biochim Acta 1986 451363–1374[Medline]
  18. ; : Bode W and Huber R. Natural protein proteinase inhibitors and their interaction with proteinases. Eur J Biochem 1992 204433–451[Medline]
  19. ; : Yun-Ge Zhao A. Expression of matrix metalloproteinase –2, –9 and tissue inhibitors of metalloproteinase –1, –2, –3 mRNAs in rat uterus during early pregnancy. Mol Reprod Dev 2002 62149–158[CrossRef][Medline]
  20. ; : Rudolph-Owen LA, Hulboy DL, Wilson CL, Mudgett J, Matrisian LM. Coordinate expression of matrix metalloproteinase family members in the uterus of normal, matrilysin-deficient, and stromelysin-1-deficient mice. Endocrinology 1997 1384902–4911[Abstract/Free Full Text]
  21. ; : Fata JE, Ho AT, Leco KJ, Moorehead RA, Khokha R. Cellular turnover and extracellular matrix remodeling in female reproductive tissues: functions of metalloproteinases and their inhibitors. Cell Mol Life Sci 2000 5777–95[CrossRef][Medline]
  22. ; : Curry TE Jr and Osteen KG. Cyclic changes in the matrix metalloproteinase system in the ovary and uterus. Biol Reprod 2001 641285–1296[Abstract/Free Full Text]
  23. ; : Curry TE Jr and Osteen KG. The matrix metalloproteinase system: changes, regulation, and impact throughout the ovarian and uterine reproductive cycle. Endocr Rev 2003 24428–465[Abstract/Free Full Text]
  24. ; : Lorenzl S, Albers DS, Narr S, Chirichigno J, Beal MF. Expression of MMP-2, MMP9, and MMP-1 and their endogenous counterregulators TIMP-1 and TIMP-2 in postmortem brain tissue of Parkinson's disease. Exp Neurol 2002 17813–20[CrossRef][Medline]
  25. ; : Olson MW, Gervasi DC, Mobashery S, Fridman R. Kinetic analysis of the binding of human matrix metalloproteinase-2 and –9 to tissue inhibitor of metalloproteinase (TIMP)-1 and TIMP-2. J Biol Chem 1997 27229975–29983[Abstract/Free Full Text]
  26. ; : Stefansson S, Aimes RT, Seward NB, Alexander DS, Quigley JP. Native TIMP-free 70 kDa progelatinase (MMP-2) secreted at elevated levels by RSV transformed fibroblasts. J Cell Physiol 1994 161419–428[CrossRef][Medline]
  27. ; : Kolkenbrock H, Hecker-Kia A, Orgel D, Ulbrich N, Will H. Activation of progelatinase A and progelatinase A/TIMP-2 complex by membrane type 2-matrix metalloproteinase. Biol Chem 1997 37871–76[Medline]
  28. ; : Stetler-Stevenson WG, Krutzsch HC, Liotta LA. Tissue inhibitor of metalloproteinase (TIMP-2). A new member of the metalloproteinase inhibitor family. J Biol Chem 1989 26417374–17378[Abstract/Free Full Text]
  29. ; : Das S, Mandal M, Chakraborti T, Mandal A, Chakraborti S. Isolation of MMP-2 from MMP-2/TIMP-2 complex: characterization of the complex and the free enzyme in pulmonary vascular smooth muscle plasma membrane. Biochim Biophys Acta 2004 1674158–174[Medline]
  30. ; : Twining SS, Schulte DP, Zhou X, Wilson PM, Fish BL, Moulder JE. Changes in rat corneal matrix metalloproteinases and serine proteinases under vitamin A deficiency. Curr Eye Res 1997 16158–165[CrossRef][Medline]
  31. ; : Nam TJ, Busby WH Jr, Clemmons DR. Characterization and determination of the relative abundance of two types of insulin-like growth factor binding protein-5 proteases that are secreted by human fibroblasts. Endocrinology 1996 1375530–5536[Abstract]
  32. ; : Dubin G. Proteinaceous cysteine protease inhibitors. Cell Mol Life Sci 2005 62653–669[CrossRef][Medline]
  33. ; : Chen B, Wen Y, Yu X, Polan ML. Elastin metabolism in pelvic tissues: is it modulated by reproductive hormones? Am J Obstet Gynecol 2005 1921605–1613[CrossRef][Medline]
  34. ; : Draper DL, Landers DV, Krohn MA, Hillier SL, Wiesenfeld HC, Heine RP. Levels of vaginal secretory leukocyte protease inhibitor are decreased in women with lower reproductive tract infections. Am J Obstet Gynecol 2000 1831243–1248[CrossRef][Medline]
  35. ; : Rinne A, Kirschke H, Jarvinen M, Hopsu-Havu VK, Wiederanders B, Bohley P. Localization of cathepsin H and its inhibitor in the skin and other stratified epithelia. Arch Dermatol Res 1985 277190–194[CrossRef][Medline]
  36. ; : Dickinson DP. Cysteine peptidases of mammals: their biological roles and potential effects in the oral cavity and other tissues in health and disease. Crit Rev Oral Biol Med 2002 13238–275[Abstract/Free Full Text]
  37. ; : Kos J, Werle B, Lah T, Brunner N. Cysteine proteinases and their inhibitors in extracellular fluids: markers for diagnosis and prognosis in cancer. Int J Biol Markers 2000 1584–89[Medline]
  38. ; : Slayden OD, Hettrich K, Carroll RS, Otto LN, Clark AL, Brenner RM. Estrogen enhances cystatin C expression in the macaque vagina. J Clin Endocrinol Metab 2004 89883–891[Abstract/Free Full Text]
  39. ; : Chughtai B and O'Riordan TG. Potential role of inhibitors of neutrophil elastase in treating diseases of the airway. J Aerosol Med 2004 17289–298[CrossRef][Medline]
  40. ; : Forsyth S, Horvath A, Coughlin P. A review and comparison of the murine alpha1-antitrypsin and alpha1-antichymotrypsin multigene clusters with the human clade A serpins. Genomics 2003 81336–345[CrossRef][Medline]



This article has been cited by other articles:


Home page
 Biol. Reprod.Home page
C. K. Wieslander, D. D. Rahn, D. D. McIntire, J. F. Acevedo, P. G. Drewes, H. Yanagisawa, and R. A. Word
Quantification of Pelvic Organ Prolapse in Mice: Vaginal Protease Activity Precedes Increased MOPQ Scores in Fibulin 5 Knockout Mice
Biol Reprod, March 1, 2009; 80(3): 407 - 414.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Pathol.Home page
D. D. Rahn, J. F. Acevedo, S. Roshanravan, P. W. Keller, E. C. Davis, L. Y. Marmorstein, and R. A. Word
Failure of Pelvic Organ Support in Mice Deficient In Fibulin-3
Am. J. Pathol., January 1, 2009; 174(1): 206 - 215.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
78/3/521    most recent
biolreprod.107.063024v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow My Folders
Right arrow Download to citation manager
Right arrow reprints & permissions
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.
Agricola
Right arrow Articles by Wieslander, C. K.
Right arrow Articles by Word, R. A.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS