Abstract
Bronchial hyperresponsiveness in mild to moderate asthma may result from airway smooth muscle cell proliferation or acquisition of a hypercontractile phenotype. Because these cells have not been well characterized in mild to moderate asthma, we examined the morphometric and gene expression characteristics of smooth muscle cells in this subgroup of patients with asthma. Using bronchial biopsies from 14 subjects with mild to moderate asthma and 15 control subjects, we quantified smooth muscle cell morphology by stereology and the expression of a panel of genes related to a hypercontractile phenotype of airway smooth muscle, using laser microdissection and two-step real-time polymerase chain reaction. We found that airway smooth muscle cell size was similar in both groups, but cell number was nearly twofold higher in subjects with asthma (p = 0.03), and the amount of smooth muscle in the submucosa was increased 50–83% (p < 0.005). Gene expression profiling in smooth muscle cells showed no difference in the expression of genes encoding phenotypic markers in cells from healthy subjects and subjects with asthma (all p > 0.1). We conclude that airway smooth muscle proliferation is a pathologic characteristic of subjects with mild to moderate asthma. However, smooth muscle cells in mild to moderate asthma do not show hypertrophy or gene expression changes of a hypercontractile phenotype observed in vitro.
Bronchial hyperresponsiveness and variable airway obstruction are cardinal features of asthma, and abnormalities of airway smooth muscle have been considered important contributors to these pathophysiologic changes (1, 2). Possible mechanisms include (1) mechanical consequences of increased amount of smooth muscle in the airway submucosa, (2) increased expression of contractile proteins in smooth muscle cells, and (3) increased activation of the contractile apparatus.
Although mathematical models suggest that an increased amount of smooth muscle in the airway submucosa is sufficient to cause bronchial hyperresponsiveness through mechanical effects alone (3–5), in vitro studies have suggested that smooth muscle cells have phenotypic plasticity (6, 7) and can manifest structural changes associated with increased intrinsic contractility. For example, under conditions of serum deprivation, a subset of dog tracheal smooth muscle cells manifests a hypercontractile phenotype characterized by increased cell length and by increased expression of a set of smooth muscle–specific contractile proteins (e.g., α-actin, smooth muscle myosin heavy chain isoforms, SM22, and myosin light chain kinase) (8, 9). One study reported increased expression of myosin light chain kinase in bronchial biopsies from human subjects with asthma, using semiquantitative reverse transcription-polymerase chain reaction (RT-PCR) (10). However, quantitative expression profiling of phenotypic markers in isolated smooth muscle cells from individuals with asthma has not otherwise been reported. Although an increased amount of smooth muscle in the airway submucosa (11, 12) and smooth muscle cell hypertrophy have been reported in subjects with severe asthma (12, 13), smooth muscle remodeling has not been established in mild to moderate asthma. These patients constitute the majority of patients with asthma and, despite having well-preserved lung function, nonetheless have significant bronchial hyperresponsiveness.
We hypothesized that the mechanism of bronchial hyperresponsiveness in mild to moderate asthma may involve structural changes to airway smooth muscle cells or adoption of a hypercontractile phenotype by these cells. To test these hypotheses we examined morphometric and gene expression characteristics of smooth muscle cells in bronchial biopsies from asthmatic subjects and healthy control subjects. We used design-based stereology to quantify morphometric characteristics of smooth muscle cells and two-step real-time PCR of RNA from laser-dissected smooth muscle cells to quantify the expression of five genes encoding smooth muscle cell structural and contractile proteins that are considered markers of smooth muscle cell phenotype. Some of the results of these studies have been previously reported in the form of an abstract (14).
We enrolled 32 subjects (16 asthmatic and 16 nonasthmatic) in a two-visit cross-sectional study. Inclusion criteria were age, 18–55 years, with and without asthma. Subjects with asthma all had a prior physician diagnosis of asthma and a provocative concentration of methacholine causing a 20% decrease in FEV1 (PC20) of less than 8 mg/ml at Visit 1, documenting bronchial hyperresponsiveness. Subjects without asthma were not specifically excluded for a history of allergic rhinitis, but were required to have a PC20 exceeding 16 mg/ml on methacholine challenge testing. Subjects with asthma were using only inhaled β-agonist medications for therapy. No subjects were using inhaled corticosteroids or leukotriene antagonists. Subjects were excluded for a respiratory infection or asthma exacerbation within the previous 6 weeks, a significant smoking history (defined as more than 10 pack-years lifetime or any cigarette smoking in the last year), significant medical problems other than asthma, or a history of treatment in the intensive care unit or intubation for acute asthma. Written informed consent was obtained from all subjects and the study was approved by the University of California, San Francisco Committee on Human Research.
Of the 32 subjects enrolled, 2 subjects did not have paraffin-embedded biopsies for morphometric analysis. Of the remaining 30 subjects, 1 subject had only 2 biopsies, and this subject was omitted from the analysis a priori to ensure adequate attention to interbiopsy variability. Thus, 29 subjects (14 with and 15 without asthma) underwent morphometric analysis (Table 1)
Morphometry | Laser Capture Microdissection | |||||
|---|---|---|---|---|---|---|
| Asthma | Healthy | Asthma | Healthy | |||
| n | 14 | 15 | 11 | 8 | ||
| Age, yr | 34.4 ± 9.0 | 31.2 ± 6.7 | 36.2 ± 9.1 | 33.8 ± 6.6 | ||
| Sex, female:male | 9:5 | 7:8 | 8:3 | 4:4 | ||
| Race | ||||||
| White | 9 | 10 | 6 | 7 | ||
| Other | 5 | 5 | 5 | 1 | ||
| FEV1, % predicted | 86 ± 14 | 105 ± 13* | 88 ± 11 | 103 ± 16† | ||
| PC20, mg/ml | 0.5 (0.1, 5) | 64 (20, 64)* | 0.7 (0.2, 6.7) | 64 (20, 64)† | ||
| AST > 2 | 11 (79%) | 6 (40%)‡ | 9 (82%) | 3 (38%)§ | ||
Procedures at Visit 1 included medical history and physical examination, an asthma characterization questionnaire, spirometry, and methacholine challenge testing. At Visit 2, bronchoscopy was performed after pretreatment with 180 μg of albuterol for all subjects (patients with asthma and healthy control subjects), using specific methods previously described (15).
Spirometry was performed according to American Thoracic Society criteria (16), using a dry rolling seal spirometer (model VRS2000; PDS Instrumentation/S&M Instrument, Doylestown, PA). Methacholine challenge was performed as previously described (17).
During bronchoscopy, 6–10 bronchial biopsies were obtained from second through fourth order carina, using spiked, fenestrated forceps (Pentax KH2411S and Pentax 8228; Pentax Precision Instrument Corporation, Orangeburg, NY). When 10 biopsies were available, 6 were reserved for morphometric analysis and 4 were immediately frozen in tissue embedding medium (Triangle Biomedical Sciences, Durham, NC) on crushed dry ice in the bronchoscopy suite.
Biopsies for morphometric analysis were formalin-fixed and paraffin-embedded as described previously (18). Each biopsy was then individually embedded in a 4-mm spherical mold to introduce isotropic uniform random orientation (19). The six spherical molds from each subject were then reembedded together in a single conventional mold and sections were cut to a thickness of 3 μm. The smooth muscle volume fraction was measured in sections stained for α-smooth muscle actin (clone 1A4; NeoMarkers, Fremont, CA) (Figure 1A)
Figure 1. Stereology and laser capture microdissection of airway smooth muscle from bronchial biopsies. (A and B) Preparation of bronchial biopsies for stereology. (A) α-Smooth muscle actin-immunostained, paraffin-embedded section. (B) Gomori's trichrome-stained, paraffin-embedded section. (C–I) Laser capture microdissection. (C) Hematoxylin and eosin-stained frozen section. A pulsed ultraviolet laser beam (VSL-337ND-S nitrogen laser; Laser Science, Franklin, MA) was focused through the objective of an inverted microscope (Axiovert S100; Zeiss, Oberkochen, Germany) to a beam spot less than 1 μm in diameter. At ×870 total magnification, cells of interest were outlined and the tissue and membrane were circumferentially dissected for nearly 360°, leaving a small attachment of membrane and tissue, termed a “bridge” (D and G). (E and H) The isolated specimen was ejected from the slide by firing the laser directly at the bridge and catapulting the cells into an inverted microcentrifuge tube cap suspended over the section and coated with PCR-grade mineral oil (Sigma). (F and I) Successful catapulting was documented by direct visualization in the Eppendorf cap.
[More] [Minimize]Frozen sections were cut to a thickness of 5 μm, mounted onto polyethylene naphthol membrane-coated glass slides (PALM Mikrolaser Technologies, Bernried, Germany), and ethanol fixed for 30 seconds (100 μl of 70% ethanol was pipetted directly onto the section and removed by suction). Using the same technique, the section was stained with Mayer's hematoxylin (Sigma) and eosin Y with phloxine B (Sigma), dehydrated in graded alcohols (70, 95, and 100%), and air dried quickly with forced air from a canister (Air Blast; InterAct Accessories, Lake Mary, FL). Laser capture microdissection was performed immediately. For these samples, the time from sectioning to completion of laser capture microdissection did not exceed 20 minutes.
Smooth muscle and epithelial cells were microdissected with the PALM laser microbeam system (PALM Mikrolaser Technologies) (Figures 1C–1I). After capture of one to six fields per section, the tissues were lysed in RNA isolation buffer (RLT buffer; Qiagen, Valencia, CA) with linear acrylamide (20 μg/ml; Ambion, Austin, TX) and samples were stored at –80°C until RNA isolation. RNA was isolated according to the RNeasy minikit protocol (Qiagen).
Gene expression in laser-captured smooth muscle and epithelial cells was measured by a two-step real-time PCR method described in detail previously (22). This method employs two separate nested sets of gene-specific primers designed to produce amplicons smaller than 250 bp. The outflanking set of primers is used in multiplex PCR amplification. The second nested set is used with a TaqMan probe for real-time PCR gene quantification on the resulting cDNA product. Therefore, this two-step method provides additional levels of gene specificity as compared with conventional RT-PCR. Real-time PCR has additional advantages over conventional RT-PCR in that it ensures measurement during the geometric rather than exponential phase of PCR when the reaction becomes limited by the substrate, and may not reflect the original representation of gene transcripts. Furthermore, this method allows normalization to the expression of housekeeping genes quantified concurrently. For this study, primer and probe sets for genes of interest were designed with Primer Express software (Applied Biosystems, Foster City, CA) on the basis of sequencing data from National Center for Biotechnology Information databases and purchased from Biosearch Technologies (Novato, CA). The primer and probe sequences employed are available for review at http://asthmagenomics.ucsf.edu/pubs/publications.htm. In the case of caldesmon, two primer and probe sets were developed to identify isoform variants generated from the same gene by RNA-splicing events. One set identifies all the isoforms, whereas caldesmon (variant 1) identifies the longest isoform variant. Total RNA was reverse transcribed with random hexamers and the RT product was amplified with gene-specific primers and multiplex hot start PCR for 25 cycles. Transcript quantifications were run with –RT cDNA controls on an ABI Prism 7700 sequence detection system (Applied Biosystems), using gene-specific primers and probes yielding cycle threshold values for each gene. Cycle thresholds were then converted into relative transcript copy number through logarithmic transformation and the application of a transformation factor based on linear regression of prior data as previously described (22). Transcript copy numbers were normalized using a two-step approach. First, the amount of starting cDNA used in TaqMan profiling was normalized on the basis of housekeeping gene expression. Then, a panel of 10 housekeeping genes was measured during TaqMan profiling, and the geometric mean value of the housekeeping genes most stably expressed across the samples was used for normalization according to Vandesompele and coworkers (23).
Measurements of airway smooth muscle volume fraction (referenced to volume of submucosa and to surface area of basal lamina) and mean airway smooth muscle cell size for asthmatic and healthy subjects were compared by unpaired t test, and measurements of number of smooth muscle cells per surface area of basal lamina were compared by Wilcoxon rank-sum test because of nonnormal distribution of these data. Correlations of morphometric measures with the forced expiratory volume in the first second (FEV1), expressed as a percentage of the predicted value, and log PC20 were performed using the Pearson correlation. Gene transcript copy numbers in asthmatic and healthy subjects were compared by Student unpaired t-test. Data analysis was performed with STATA 5.0 (StataCorp, College Station, TX).
The volume fraction of airway smooth muscle normalized to volume of submucosa (Vv smmu,su) in subjects with asthma was 50% greater than in healthy subjects (p = 0.005; Figure 2)
Figure 2. Airway smooth muscle content in the submucosa is increased in subjects with mild to moderate asthma as compared with nonasthmatic control subjects. Morphometric outcomes were calculated from point and line intersect counts as follows: Vv smmu, su = (Σ points on smooth muscle × a/p × h)/(Σ points on submucosa × a/p × h); Sv bala, su = (2 × Σ line intersections with basal lamina)/(l/p × Σ points on submucosa); Vs smmu, bala = Vv smmu, su/Sv bala, su; where a/p denotes area per point (mm2), l/p denotes length per point (mm), and h denotes the thickness of the tissue section (μm). Results for Vv smmu, su with means ± SD are presented here.
[More] [Minimize]Mean smooth muscle cell size was similar in healthy and asthmatic subjects (2,654 ± 757 versus 2,911 ± 634 μm3/cell, respectively; p = 0.33) (Figure 3)
Figure 3. Mean airway smooth muscle cell volume is normal in mild to moderate asthma, but the number of cells is increased. Mean airway smooth muscle cell volume was calculated from the number of cells per volume of smooth muscle surveyed. The number of cells per surface area of basal lamina was calculated on the basis of the number of cells per volume of smooth muscle surveyed and the volume of smooth muscle per surface area of basal lamina previously calculated for each subject. *p = 0.03.
[More] [Minimize]To establish the specificity of laser capture, gene expression for muscle-specific and epithelial cell-specific genes was measured in smooth muscle cells and in epithelial cells dissected from the same section in three subjects with asthma and three healthy subjects (Table 2)
Transcript Number × 106 | |||
|---|---|---|---|
| Gene | Smooth Muscle | Epithelial | |
| Smooth muscle–specific | |||
| α-Smooth muscle actin | 1,270 ± 372 | 4.2 ± 1.9 | |
| Caldesmon | 286 ± 137 | 1.1 ± 1.9 | |
| Caldesmon V1 | 120 ± 37 | 0.5 ± 1.0 | |
| Calponin | 267 ± 83 | < 0.001 | |
| Smooth muscle myosin heavy chain (SM1) | 686 ± 135 | 1.4 ± 1.3 | |
| SM22a | 2,370 ± 768 | 17.1 ± 13 | |
| Myosin light chain kinase | 104 ± 40 | 5.1 ± 4.2 | |
| Epithelial cell–specific | |||
| Muc2 | < 0.001 | 3.7 ± 3.7 | |
| Muc5ac | < 0.001 | 321 ± 471 | |
| Muc5b | < 0.001 | 38 ± 32.7 | |
| Trefoil factor 3 | < 0.001 | 106 ± 172 | |
| Aquaporin 5 | < 0.001 | 70 ± 53 | |
| CFTR | 0.002 | 5.8 ± 5.2 | |
To establish the variability in gene expression profiling attributable to laser capture and processing, the same bundle of smooth muscle was captured in three successive sections from a single biopsy. The mean coefficient of variation for transcript copy number across a panel of 48 genes examined was 0.49. This variability was significantly less than the mean variability observed between different biopsies from the same subject (mean coefficient of variation, 0.71), which was, in turn, less than the variability observed between individuals (mean coefficient of variation, 0.97).
When we tested our hypothesis that the expression of genes encoding structural and contractile proteins, as markers of a hypercontractile phenotype, would be upregulated in smooth muscle from subjects with asthma, we found that the expression levels of these genes was not significantly different from normal (all comparisons, p > 0.10; Figure 4)
Figure 4. Expression of genes encoding markers of a hypercontractile airway smooth muscle cell phenotype is normal in mild to moderate asthma. Expression levels were measured by laser capture microdissection and two-step real-time PCR. Data labeled caldesmon were generated with a primer and probe set that identifies all caldesmon isoforms, whereas caldesmon v1 identifies the longest isoform variant. MLCK = myosin light chain kinase; SM-MHC = smooth muscle myosin heavy chain.
[More] [Minimize]We found that the number of airway smooth muscle cells in bronchial biopsies from subjects with mild to moderate asthma is nearly twofold higher than normal, leading to an overall increase in the volume fraction of smooth muscle in asthma of 50–83%. These findings are consistent with reports that asthmatic smooth muscle cells demonstrate increased proliferative capacity in vitro (24), and provide a rationale for research efforts to identify specific mediators of increased proliferation of airway smooth muscle in mild to moderate asthma. Because mathematical models suggest that increasing the volume of smooth muscle in the airway can lead to bronchial hyperresponsiveness through local mechanical effects (3–5), mediators of smooth muscle cell hyperplasia would be logical targets for novel therapies for bronchial hyperresponsiveness in mild to moderate asthma.
Applying laser capture microdissection to isolate airway smooth muscle bundles from biopsy tissue sections, we were able to measure gene expression in airway smooth muscle, using TaqMan-based gene-profiling methods. We measured the expression profile of a panel of five genes (α-actin, smooth muscle myosin heavy chain isoforms, SM22, and myosin light chain kinase) considered markers of a hypercontractile phenotype, and we found that these genes were not upregulated in asthmatic smooth muscle. Thus, we were unable to find evidence of phenotypic change of airway smooth muscle in mild to moderate asthma at the gene expression level. Phenotypic change of airway smooth muscle has been observed in vitro and in an animal model of airway inflammation (7–9, 25). For example, under conditions of serum deprivation, a subset of dog tracheal airway smooth muscle cells in culture manifests a hypercontractile phenotype characterized by increased cell length, and increased expression of α-actin, smooth muscle myosin heavy chain isoforms, SM22, and myosin light chain kinase (8, 9). Alternatively, airway smooth muscle cells may dedifferentiate and acquire features of a synthetic/proliferative cell with decreased expression of structural and contractile proteins (7, 25). This phenotypic change occurs in a wide range of smooth muscle cells, including dog, rat, and human, and represents another form of phenotypic change potentially important in asthma. With the application of laser capture microdissection and two-step real-time PCR, we found that the expression profiles of genes encoding α-actin, smooth muscle myosin heavy chain isoforms, SM22, and myosin light chain kinase were similar in patients with mild to moderate asthma and in healthy subjects. It remains possible that phenotypic plasticity may occur through a change in structural and contractile protein content over time without a discernible change in expression of the genes encoding these proteins at any given time point, or that gene expression changes are apparent only during periods of asthma exacerbation. The current study cannot address the latter possibility directly because subjects were excluded for a recent exacerbation. However, methods of protein analysis are not currently applicable to the extremely small samples obtained by laser capture microdissection. It also is possible that we are not able to detect 25–30% differences in gene expression with the currently available application of laser capture microdissection and that this could cause us to miss potentially clinically important differences. Finally, it is also possible that phenotypic change may occur at other locations in the airway or in more severe disease.
We took several steps to protect morphometric measures from potential sources of bias. Most importantly, we applied methods from design-based stereology, a discipline that incorporates systematic random sampling and accounts for the three-dimensional nature of tissues and cells (21). Furthermore, we excluded adjacent connective tissue from measurements by using both a smooth muscle-specific immunostain (α-smooth muscle actin) and a collagen-specific stain (Gomori's trichrome), and making measurements at relatively high magnification (×870, total). This approach was necessary because previous findings of increased airway smooth muscle content in severe asthma have been criticized as potentially biased by the inclusion of connective tissue (26, 27).
Our data indicating no increase in myosin light chain kinase expression in asthma contrast with the data of Ma and coworkers (10), who found increased gene expression for myosin light chain kinase in bronchial biopsies from subjects with asthma. Several reasons could explain this difference. Whereas we analyzed laser-captured smooth muscle cells by real-time RT-PCR, Ma and coworkers (10) relied on semiquantitative RT-PCR analysis of mRNA, using either whole biopsy homogenates or smooth muscle cells manipulated ex vivo through enzymatic dispersion. Because we have demonstrated that biopsy smooth muscle content is increased in asthma, the use of homogenates poses a challenge to normalization of expression data.
As indicated above, other mechanisms besides smooth muscle cell proliferation may still contribute to smooth muscle contractility and airway hyperresponsiveness in asthma. In particular, increased activation of the contractile apparatus may occur in asthma through increased tone or increased stimulation by contractile mediators without changes in the expression of structural and contractile proteins. Indeed, physiologic measurements made in enzymatically dispersed cells demonstrate increases in maximum shortening capacity and velocity in smooth muscle cells from subjects with asthma (10). Unfortunately, physiologic abnormalities cannot be evaluated by the techniques described here. Other potential contributors to airway smooth muscle hyperresponsiveness in asthma that are poorly studied using these techniques include changes in the elastance of tissues surrounding smooth muscle cells (28). Furthermore, it is possible that phenotypic change of smooth muscle is important in disease more severe than that studied here, or in myofibroblasts that may be present outside of the smooth muscle bundle. Because we limited our microdissection to smooth muscle cells within bundles, our data do not pertain to myofibroblasts.
We conclude that there is hyperplasia of airway smooth muscle cells in mild to moderate asthma, but that cellular hypertrophy is absent and gene expression levels of contractile proteins considered markers of a hypercontractile phenotype are not increased. Taken together, these data show that smooth muscle proliferation is a characteristic of mild to moderate asthma, and that targeting smooth muscle cell proliferation may represent a strategy for treating bronchial hyperresponsiveness in asthma.
The authors thank James K. Brown, M.D., for helpful comments on the manuscript.
| 1. | Seow CY, Schellenberg RR, Pare PD. Structural and functional changes in the airway smooth muscle of asthmatic subjects. Am J Respir Crit Care Med 1998;158:S179–S186. |
| 2. | Martin JG, Duguet A, Eidelman DH. The contribution of airway smooth muscle to airway narrowing and airway hyperresponsiveness in disease. Eur Respir J 2000;16:349–354. |
| 3. | James AL, Pare PD, Hogg JC. The mechanics of airway narrowing in asthma. Am Rev Respir Dis 1989;139:242–246. |
| 4. | Lambert RK, Wiggs BR, Kuwano K, Hogg JC, Pare PD. Functional significance of increased airway smooth muscle in asthma and COPD. J Appl Physiol 1993;74:2771–2781. |
| 5. | Macklem PT. A theoretical analysis of the effect of airway smooth muscle load on airway narrowing. Am J Respir Crit Care Med 1996;153:83–89. |
| 6. | Halayko AJ, Salari H, Ma X, Stephens NL. Markers of airway smooth muscle cell phenotype. Am J Physiol 1996;270:L1040–L1051. |
| 7. | Hirst SJ, Walker TR, Chilvers ER. Phenotypic diversity and molecular mechanisms of airway smooth muscle proliferation in asthma. Eur Respir J 2000;16:159–177. |
| 8. | Halayko AJ, Camoretti-Mercado B, Forsythe SM, Vieira JE, Mitchell RW, Wylam ME, Hershenson MB, Solway J. Divergent differentiation paths in airway smooth muscle culture: induction of functionally contractile myocytes. Am J Physiol 1999;276:L197–L206. |
| 9. | Ma X, Wang Y, Stephens NL. Serum deprivation induces a unique hypercontractile phenotype of cultured smooth muscle cells. Am J Physiol 1998;274:C1206–C1214. |
| 10. | Ma X, Cheng Z, Kong H, Wang Y, Unruh H, Stephens NL, Laviolette M. Changes in biophysical and biochemical properties of single bronchial smooth muscle cells from asthmatic subjects. Am J Physiol Lung Cell Mol Physiol 2002;283:L1181–L1189. |
| 11. | James A, Carroll N. Airway smooth muscle in health and disease; methods of measurement and relation to function. Eur Respir J 2000;15:782–789. |
| 12. | Ebina M, Takahashi T, Chiba T, Motomiya M. Cellular hypertrophy and hyperplasia of airway smooth muscles underlying bronchial asthma: a 3-D morphometric study. Am Rev Respir Dis 1993;148:720–726. |
| 13. | Benayoun L, Druilhe A, Dombret MC, Aubier M, Pretolani M. Airway structural alterations selectively associated with severe asthma. Am J Respir Crit Care Med 2003;167:1360–1368. |
| 14. | Woodruff PG, Dolganov GM, Ferrando RE, Carter R, Solberg OD, Hays SR, Wong HH, Cadbury PS, Fahy JV. Mild to moderate asthma is characterized by hyperplasia of airway smooth muscle cells without hypertrophy or acquisition of a hypercontractile phenotype [abstract]. Am J Respir Crit Care Med 2003;167:A33. |
| 15. | Fahy JV, Wong H, Liu J, Boushey HA. Comparison of samples collected by sputum induction and bronchoscopy from asthmatic and healthy subjects. Am J Respir Crit Care Med 1995;152:53–58. |
| 16. | American Thoracic Society. Standardization of spirometry: 1994 update. Am J Respir Crit Care Med 1995;152:1107–1136. |
| 17. | Claman DM, Boushey HA, Liu J, Wong H, Fahy JV. Analysis of induced sputum to examine the effects of prednisone on airway inflammation in asthmatic subjects. J Allergy Clin Immunol 1994;94:861–869. |
| 18. | Hays SR, Woodruff PG, Khashayar R, Ferrando RE, Liu J, Fung P, Zhao CQ, Wong HH, Fahy JV. Allergen challenge causes inflammation but not goblet cell degranulation in asthmatic subjects. J Allergy Clin Immunol 2001;108:784–790. |
| 19. | Nyengaard JR, Gundersen HJG. The isector: a simple and direct method for generating isotropic, uniform random sections from small specimens. J Microsc 1992;165:427–431. |
| 20. | Sterio DC. The unbiased estimation of number and sizes of arbitrary particles using the disector. J Microsc 1984;134:127–136. |
| 21. | Bolender RP, Hyde DM, Dehoff RT. Lung morphometry: a new generation of tools and experiments for organ, tissue, cell, and molecular biology. Am J Physiol 1993;265:L521–L548. |
| 22. | Dolganov GM, Woodruff PG, Novikov AA, Zhang Y, Ferrando RE, Szubin R, Fahy JV. A novel method of gene transcript profiling in airway biopsy homogenates reveals increased expression of a Na+-K+-Cl– cotransporter (NKCC1) in asthmatic subjects. Genome Res 2001;11:1473–1483. |
| 23. | Vandesompele J, De Preter K, Pattyn F, Poppe B, Van Roy N, De Paepe A, Speleman F. Accurate normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes. Genome Biol 2002;3:RESEARCH0034. |
| 24. | Johnson PR, Roth M, Tamm M, Hughes M, Ge Q, King G, Burgess JK, Black JL. Airway smooth muscle cell proliferation is increased in asthma. Am J Respir Crit Care Med 2001;164:474–477. |
| 25. | Moir LM, Leung SY, Eynott PR, McVicker CG, Ward JP, Chung KF, Hirst SJ. Repeated allergen inhalation induces phenotypic modulation of smooth muscle in bronchioles of sensitized rats. Am J Physiol Lung Cell Mol Physiol 2003;284:L148–L159. |
| 26. | Thomson RJ, Schellenberg RR. Increased amount of airway smooth muscle does not account for excessive bronchoconstriction in asthma. Can Respir J 1998;5:61–62. |
| 27. | Thomson RJ, Bramley AM, Schellenberg RR. Airway muscle stereology: implications for increased shortening in asthma. Am J Respir Crit Care Med 1996;154:749–757. |
| 28. | Bramley AM, Thomson RJ, Roberts CR, Schellenberg RR. Hypothesis: excessive bronchoconstriction in asthma is due to decreased airway elastance. Eur Respir J 1994;7:337–341. |
| 29. | National Heart, Lung, and Blood Institute. Guidelines for the diagnosis and management of asthma: update on selected topics 2002. National Asthma Education and Prevention Program Expert Panel Report. Washington, DC: U.S. Government Printing Office; 2002. |
