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Senescent Expression of Genes Coding Collagens, Collagen-Degrading Metalloproteinases, and Tissue Inhibitors of Metalloproteinases in Rat Vocal Folds

Comparison With Skin and Lungs

^a Division of Otolaryngology, Department of Surgery, School of Medicine, University of Utah, Salt Lake City

Steven D. Gray, Division of Otolaryngology, School of Medicine, University of Utah, 100 North Medical Drive, Room 4500, Salt Lake City, UT 84123 E-mail: pcsgray{at}ihc.com.

Decision Editor: John A. Faulkner, PhD

	Abstract

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In humans, vocal tissue stiffness increases with age, suggesting a possible contribution of age-associated variations in vocal fold collagen turnover to voice senescence. The underlying mechanisms remain to be explored. With the use of reverse-transcriptase polymerase chain reaction (RT-PCR), collagen subtypes expressed in rat vocal folds were determined, and messenger RNA (mRNA) levels of collagens (types I, III, IV, and V), collagen-degrading proteinases (collagenase 3, gelatinase A and B), and tissue inhibitors of metalloproteinases (TIMP-1 to TIMP-4) were measured in vocal folds of neonatal, adult, and elderly rats. Collagens I, III–VIII, XV, XVII, and XVIII are abundantly expressed, whereas collagens II, IX, X, and XI are absent in rat vocal folds. Messenger RNA levels of collagens I, III, IV, and V and collagen-degrading proteinases in the vocal folds of the adult rats are significantly lower than those in the neonates. These mRNA levels show further decline in the vocal folds of the elderly rats, but only the decrease in mRNA levels of collagens I and V significantly differ from the adult levels. There are no marked age-related alterations in vocal fold levels of TIMP mRNAs, and the tissue variation in the gene expression of the aforementioned molecules is minute. Rat vocal folds display tissue-specific expression of collagen genes. Diminished gene expression for collagens and proteinases and unchanged gene expression for TIMPs indicate a slowdown in collagen turnover that may increase the cross-linking of collagen molecules. This observation may explain in part the stiffness that occurs with aging in human vocal folds.

THE exact mechanisms governing age-related voice changes have not yet been clearly determined. The human vocal fold has a laminar structure and a configuration that appear to be major determinants of its physical properties, and in turn its performance ⁽¹⁾. With age, human vocal folds become less elastic, more viscous, and stiff ⁽²⁾, which may contribute to vocal senescent characteristics ⁽³⁾⁽⁴⁾. It has been suggested that age-dependent alterations in the alignment and distribution of connective tissue fibers in the vocal folds may be among the mechanisms underlying the senescent aberration of vocal fold elasticity and stiffness ⁽⁵⁾⁽⁶⁾. The molecular mechanisms for age-associated changes in physical properties of vocal folds have not been explored.

Composition and levels of extracellular matrix (ECM) proteins are critical in maintaining the integrity, elasticity, viscosity, and stiffness of vocal folds ⁽²⁾. Collagens comprise the most prominent group of the ECM proteins in the vocal folds ⁽⁶⁾⁽⁷⁾. To elucidate the mechanisms by which the structure and function of the vocal folds senesce, it is valuable to appreciate age-dependent variations in turnover of the collagenous proteins in vocal folds. Such information could also provide windows through which therapeutic manipulations toward the treatment of the senescence-related voice abnormalities can be formulated. The present study is focused on determining collagen subtypes that express in rat vocal folds and quantifying senescent gene expression for collagens and collagen-degradation-related major proteinases in rat vocal folds.

The rat model is selected because it is a model often used in aging research; a good library of ECM genes are available; and with the use of microdissection techniques, the vocal folds can be harvested with minimal thyroarytenoid muscle contamination. Rat vocal folds are not precisely similar to human ones. A salient difference is that there is no vocal ligament. This is likely a result of the following concept: Cells subjected to mechanical forces may alter their gene expression. Because no other mammal phonates in a similar manner to humans in terms of duration and range, and because no other mammal has similar anatomy, the biomechanical forces associated with human phonation (oscillation, shear, and stress forces) are not found in any other animal model. Thus, studies of fibroblast behavior dealing with phonation using any animal model will have shortcomings. Most other animal models may be representative of some aspects of human phonation, such as cats and monkeys for neural control, and canines for peripheral nerve and physiology studies. Perhaps one of the best aspects of the rat model is that phonation is minimal, and consequently the biobehavior of rat vocal fold fibroblasts is not likely to be influenced by the biomechanical forces of phonation without the contamination of force effects. Because of this, it is our belief that the rat is a better model for aging studies of ECM regulation. Therefore, this study may focus on aspects of vocal fold fibroblast behavior related to aging unencumbered with gene expression influenced by mechanical forces. Other animal models may not have been as free from the influence of mechanical forces. Of course, future studies with comparative data from human vocal fold fibroblasts from different age groups will be very interesting, as they will demonstrate the composite product of biomechanical forces and aging effects.

	Methods

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Animals
Three groups of F344 Sprague Dawley rats, purchased from the National Institute on Aging (Bethesda, MD), six for each group and aged 1.5 ± 0.5 weeks (neonatal), 6.0 ± 0.5 months (adult), and 24 ± 0.5 months (elderly), were studied. As a way to obviate gender effects, only males were used. After they were adapted in the Animal Resource Center of University of Utah Health Science Center for 4 days, the animals were anesthetized; the vocal folds, a segment of skin covering 1 cm of the tail tip, and 100 mg of lung tissue were harvested from each rat. The tissues were immersed immediately in 200–1000 µl of ice-cold cell disruption solution, RNA STAT-60 (Tel-Test, Inc., Friendswood, TX), snap frozen with liquid nitrogen, and saved at -80°C until being further processed for extraction of total RNA.

Extraction of Total RNA
Total RNA was prepared from the frozen tissues with the use of a guanidine–phenol extraction technique ⁽⁸⁾. After thawed on ice, the tissues were finely homogenized. Extraction of total RNA from the tissue homogenate was performed according to the instructions provided by the manufacturer of the cell disruption solution. The quantity and the quality of the extracted RNA were determined by absorbency at 260 nm, and by visualizing the bands of 18S and 28S rRNA after 2 µg total RNA was separated on a 1% agarose–formaldehyde gel and stained with ethidium bromide, respectively.

Reverse-transcription (RT) of total RNA to the first strand of complementary DNA (cDNA)..-- As a way to eliminate genomic DNA contamination, RNA samples were treated with RNase-free Dnase I (Amersham Pharmacia Biotech, Piscataway, NJ) before being reverse-transcribed to first strand cDNA. The First Strand cDNA Synthesis Kit from Boehringer Mannheim (Indianapolis, IN) was used with a modification of the manufacturer's protocol to achieve maximum yield. The total volume of the RT reaction was 20 µl and the final concentrations of the reagents in the reaction mixture were RT buffer (1x), MgCl₂ (5 mM), deoxynucleoside triphosphate (dNTP) mixture (1 mM for each dNTP), random primers (total 0.08 A₂₆₀ units), RNase inhibitor (2.5 units/µl), and Arian myeloblastosis virus (AMV) reverse transcriptase (2.5 units/µl). The reactions were incubated in a thermal cycler (GeneAmp PCR System 2400 or 9600, Perkin Elmer, Norwalk, CA) at 25°C for 10 minutes, 42°C for 95 minutes, 99°C for 5 minutes; and 4°C for 5 minutes, consecutively. The total volume of the reaction was then brought to 400 µl with diethyl pyrocarbonate (DEPC) treated H₂O, aliquoted, and saved at -80°C until use.

Amplification of cDNA with polymerase chain reaction (PCR)..-- "Hot start PCR" with an application of an anti-Taq DNA polymerase antibody, TaqStart antibody (Clontech Lab. Inc., Palo Alto, CA), was used. The total volume of the PCR reaction was 25 µl in 1x PCR buffer (10 mM of Tris-HCl, 50 mM of KCl, pH 8.3). The optimal concentrations of the reagents (especially MgCl₂) in the PCR reactions varied with different target genes, but for PCR amplifying most target genes, a standard recipe was feasible. The final concentrations of the reagents in this standard recipe were dNTP (0.2 mM for each dNTP), MgCl₂ (1.5 mM), forward and reverse primers (0.5 µM for each), a 1:1 mixture of Taq polymerase and TaqStart antibody (0.4 µl), and various amount of cDNA equivalent to 1.25–10 ng of total RNA. For most target genes the mixture was incubated in Perkin Elmer thermal cyclers with the following standard protocol: One cycle of 94°C for 1 minute, followed by 35 cycles of 94°C for 30 seconds (denaturing), 56°C for 1 minute (annealing), and 72°C for 2 minutes (extension), and then by one cycle of 72°C for 5 minutes and cooled to 4°C.

Experimental conditions for PCR were optimized. Table 1 lists the optimal concentrations of MgCl₂ and the annealing temperature (AT) for the primers that have been used in this study. Linearity of the PCR reactions was defined as the linear relationship between the amount of the starting cDNA that was added to the PCR reactions and the amount of the corresponding PCR products. This linearity was determined, for each gene and tissue, by adding different amounts of the starting cDNA to the PCR reactions and amplifying all reactions with the same PCR conditions. The densitometric readings of the PCR products were plotted against the amounts of the starting cDNA. The optimal amount of starting cDNA to be used in formal experiments was then determined as the one that gave a linear trajectory before the linear regression line reached the maximum plateau.

For verification that a PCR product was not contamination-brought over fakery, a negative control reaction, in which cDNA was omitted, was always amplified simultaneously with the reaction for the target gene. The fidelity of the PCR amplification was confirmed by sequencing the PCR product. All the primers used in this study were initially designed according to the published gene sequences, tested previously and listed in Table 1 . The primers were synthesized and crude purified commercially.

Data Analysis
Statistical analyses were performed by using GraphPad PRISM (Version 2.0, Intuitive Software for Science, San Diego, CA). Results are expressed as the mean ± standard error (SE). Statistical significance, set at p < .05, was determined by using a one-way analysis of variance (ANOVA). Each data point represents a minimum of six individual RT-PCR assays.

	Results

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Collagen Subtypes Expressed in Rat Vocal Folds
As shown in Fig. 1, collagen types I, III–VIII, XV, XVII, and XVIII are abundantly expressed in rat vocal folds. No expression of genes coding for collagen types II, IX, X, and XI is detected in the vocal folds. As a way to ensure that the negativity was not due to inadequate amplification, PCR cycles were increased to 40, and the same results were obtained. In contrast, strong signals of collagen types II, IX, X, and XI can be detected when cartilage RNA was used as control (data not shown). Other collagen subtypes were not examined for their possible expression in the vocal folds because of a lack of information on rat genes.

View larger version (50K): [in this window] [in a new window]   Figure 1. Collagen subtypes expressed in rat vocal folds as determined by reverse-transcriptase polymerase chain reaction (RT-PCR). Total RNA was extracted from vocal folds of neonatal rats and analyzed with RT-PCR by using the primers listed in Table 1 according to the procedures as described in Materials and Methods. M = Molecular markers (50-bp DNA ladder), sizes of which are numbered on the left side of the gel. Rows 1–9 represent collagen types I, III, IV, V, VI, VII, VIII, XV, XVII, and XVIII, respectively. The expected molecular sizes of the PCR products are listed in Table 1 .

View larger version (63K): [in this window] [in a new window]   Figure 2. Expression of genes for collagen types I, III, IV and V in the vocal folds of rats of different ages as determined by reverse-transcriptase polymerase chain reaction (RT-PCR). A: Representative photogram of ethidium bromide stained agarose gel for PCR product separation. I = rats aged 1–2 weeks; II = rats aged 5–6 months; III = rats aged 24–25 months. M = molecular markers (50-bp DNA ladder), sizes of which are numbered on the leftmost of the gel. Locations of the PCR products are marked and their expected molecular sizes are listed in Table 1 . B: Bar graph results are presented as mean ± SE. *p < .05 vs the levels of rats aged 1–2 weeks; #p < .05 vs the levels in rats aged 5–6 months.

View larger version (64K): [in this window] [in a new window]   Figure 3. Expression of genes for collagenase 3, gelatinase A, and gelatinase B in the vocal folds of rats of different ages. A: Representative photogram of ethidium bromide stained agarose gel for polymerase chain reaction product separation. I = rats aged 1–2 weeks; II = rats aged 5–6 months; III = rats aged 24–25 months. M = molecular markers (50-bp DNA ladder), sizes of which are numbered on the left side of the gel. B: Bar graph results are presented as mean ± SE. *p < .05 vs the levels of rats aged 1–2 weeks.

View larger version (66K): [in this window] [in a new window]   Figure 4. Expression of genes for tissue inhibitors of metalloproteinases (TIMPs) in the vocal folds of rats of different ages. A: Representative photogram of ethidium bromide stained agarose gel for polymerase chain reaction product separation. I = rats aged 1–2 weeks; II = rats aged 5–6 months; III = rats aged 24–25 months; M = molecular markers (50-bp DNA ladder), sizes of which are numbered on the left side of the gel. B: Bar graph results are presented as mean ± SE. *p < .05 vs the levels of rats aged 1–2 weeks; #p < .05 vs the levels in rats aged 5–6 months.

View larger version (17K): [in this window] [in a new window]   Figure 5. Comparison of expression of collagen genes in vocal folds (V), tail skin (S), and lungs (L) of rats of different ages. Each data point is the mean of six PCR assays ± SE. *p < .05 vs levels in rats aged 1–2 weeks; #p < .05 vs levels in rats aged 5–6 months.

View larger version (19K): [in this window] [in a new window]   Figure 6. Comparison of expression of collagenase 3, gelatinase A, and gelatinase B in vocal folds (V), tail skin (S), and lungs (L) of rats of different ages. Each data point is the mean of six PCR assays ± SE. *p < .05 vs levels in rats aged 1–2 weeks; #p < .05 vs levels in rats aged 5–6 months.

View larger version (24K): [in this window] [in a new window]   Figure 7. Comparison of expression of TIMP-1, TIMP-2, TIMP-3, and TIMP-4 in vocal folds (V), tail skin (S), and lungs (L) of rats of different ages. Each data point is the mean of six PCR assays ± SE. *p < .05 vs levels in rats aged 1–2 weeks; #p < .05 vs levels in rats aged 5–6 months.

	Discussion

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Collagens IV and VII are basement-membrane-associated collagens ⁽⁹⁾. Collagen type IV forms the meshwork scaffold of the basement membranes, whereas the type VII is an anchoring molecule that locates in the subbasal lamina and functions to anchor the basement-membrane zone to the underlying stroma. In the human and canine vocal folds, this collagen also mainly distributes in the superficial layer of the lamina propria close to the basement membrane and organizes in such a way as to facilitate fixing the basement membrane of the vocal epithelium to the underlying connective tissue, as demonstrated by electron microscopy ⁽¹⁰⁾. In many tissues, collagens IV and VII are produced by the epithelial cells overlying basement membrane, but their source in the vocal folds remains to be found.

Collagen VIII is an endothelial-cell-associated collagen. This collagen is expressed in the vocal folds in a relatively small amount. As in other tissues, collagen VIII may be produced from capillaries present in the lamina propria of the vocal folds. The exact functions of collagen VIII, however, are largely unknown. The other collagens that are demonstrated in the vocal folds, including XV, XVII, and XVIII, are recently cloned and information on their physiological function is lacking.

In this study, collagen types II, IX, X, and XI were not detected in rat vocal folds (data not shown). The fact that these collagens are mainly associated with cartilaginous tissues ⁽⁹⁾ may be why they are absent in cartilage-lacking vocal folds. There are other recently characterized collagens that were not examined in this study. It is possible that some of them may express in the vocal folds and may even play an important role in phonation physiology.

Age-Associated Alterations in Gene Expression
These alterations are of collagens, collagen-degrading MMPs, and TIMPs in vocal folds. It is clear from our data that the gene expression of collagen types I, III, IV, and V displays an age-dependent reduction (Fig. 2). We also found that both skin and lung exhibit similar age-related changes in the gene expression of these collagen genes. Thus, an age-associated decrease in expression of these collagen genes in the vocal folds may reflect an age-dependent degenerative process that involves most systems. The results do not exclude, however, the possibility that this similarity may not exist in disease settings. This is because senescence is a process regulated both genetically and environmentally, and disease-associated extrinsic factors may affect organs or tissues differentially.

The composition and levels of ECM components are the reflection of the balance between the biosynthesis and degradation of ECM proteins. ECM turnover, either physiological or pathological, is in most cases a highly organized process that involves the selective action of a group of zinc- and calcium-dependent proteases, namely the MMPs or matrixin protease family ⁽¹¹⁾. In addition, in vivo activities of MMPs are controlled at several levels, including their interactions with specific naturally occurring inhibitors, for example, the TIMPs ⁽¹¹⁾. TIMPs are cell-secreted nonspecific inhibitors that act as negative regulators of MMPs, and four have been cloned and well characterized, designated as TIMP-1, TIMP-2, TIMP-3, and TIMP-4, respectively. An imbalance between MMPs and TIMPs results in either activation or suppression of MMPs, and in turn determines the rate of matrix accumulation and degradation. In rat vocal folds, gene expression of collagen-degrading MMPs appears to adopt a pattern that is different from the one of TIMPs. As compared with that of neonatal rats, vocal levels of mRNAs for MMPs significantly diminish in adult rats and do not show a further decrease in elderly rats (Fig. 3). In contrast, although gene expression of TIMP-1, TIMP-3, and TIMP-4 in the vocal folds remains at stable levels at different ages, the mRNA level of TIMP-2 in elderly rats significantly declines to 58% of the neonatal level (Fig. 4). These data indicate that a collagen degradative process in vocal folds becomes less active from adulthood on as a result of the downregulation in gene expression of collagen-degrading MMPs and may be further enhanced in aged vocal folds because of an increased expression of some TIMP genes.

Conclusions
It is well observed clinically that stiffness of the human vocal fold lamina propria increases with age ⁽³⁾⁽⁴⁾. One theory holds that this is due to age-dependent alterations in alignment and distribution of connective tissue fibers in the vocal folds ^(5)(6)(7). Our data demonstrate that in aging rats, without the influence of phonatory forces, the balance between production and degradation of vocal collagens may lead to a diminished collagen volume. In contrast, in humans, quantitative histology studies observe either an unchanged or an increased collagen volume in the vocal folds of the elderly population ^(5)(6)(7). We speculate that, as in other organs, there may be an enhanced cross-linking of collagen molecules in aged vocal folds, which produces over-cross-linked collagen fibers that are resistive to digestive enzymes ^(12)(13) as well as to deformation ⁽¹⁴⁾. One physical reflection of these subtle molecular changes is the increase in vocal stiffness. Another possibility for the increase in collagen fibers seen in humans is that the biomechanical forces from phonation influence collagen gene expression. It is possible that the effects of biomechanical forces account for the increase in collagen seen in elderly human vocal folds, whereas the aging effects of decreased collagen turnover and likely increased collagen cross-linking demonstrated in this rat model explain the stiffness found in elderly human folds. Further experiments are necessary to confirm this hypothesis.

	Acknowledgments

 
This work was supported by National Institutes of Health Grant R01 DC04336-01. We thank Mindy Simon for her assistance in manuscript preparation.

Received March 24, 2000

Accepted July 18, 2000

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