Journal of Toxicology and Environmental Health
ISSN: 0098-4108 (Print) (Online) Journal homepage: http://www.tandfonline.com/loi/uteh19
Ciliated respiratory epithelial surface changes after formaldehyde exposure Frank Colizzo , Mori J. Krantz , James E. Fish & Annette T. Hastie To cite this article: Frank Colizzo , Mori J. Krantz , James E. Fish & Annette T. Hastie (1992) Ciliated respiratory epithelial surface changes after formaldehyde exposure, Journal of Toxicology and Environmental Health, 35:4, 221-234, DOI: 10.1080/15287399209531613 To link to this article: http://dx.doi.org/10.1080/15287399209531613
Published online: 20 Oct 2009.
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Date: 07 November 2015, At: 22:54
CILIATED RESPIRATORY EPITHELIAL SURFACE CHANGES AFTER FORMALDEHYDE EXPOSURE Frank Colizzo, Mori J. Krantz, James E. Fish, Annette T. Hastie
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Department of Medicine, Thomas Jefferson University, Philadelphia, Pennsylvania
The investigation sought to identify alterations of specific ciliated epithelial surface components after exposure to formaldehyde (HCHO) levels that decrease respiratory ciliary function. Bovine tracheae were reacted with an analog of N-hydroxysuccinimidobiotin to label epithelial surface-accessible components before exposure to HCHO. The tracheae were then exposed to 0, 16, 33, and 66 ng HCHO/cm2 epithelial surface for 30 min. Cilia were isolated from the epithelium, separated into membrane and internal axonemal portions, analyzed on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PACE), and either stained to detect proteins or transblotted to detect biotin-labeled components. Densitometric analysis of axoneme proteins showed a decrease in the total amount extracted with increased HCHO concentration, including axoneme-specific proteins, dynein, and tubulin. However, biotinylated proteins in the axoneme fractions proportionately increased. Membrane fractions showed little change in protein with increasing HCHO concentration. The majority of these is not biotin-labeled and thus not surface-accessible components. Biotinylated material in the membrane fractions showed a significant decrease with increased HCHO concentration, particularly of bands at 92, 98, and 105 kD. These data suggest that increasing HCHO exposure reduces both extractable ciliary axonemes and detergent-soluble surface components, possibly by stabilizing respiratory epithelial membranes. This process apparently strengthens association of certain surface components with the internal axoneme, thereby reducing subsequent solubilization in detergent.
INTRODUCTION Formaldehyde (HCHO) occurs widely in industrial and domestic environments, resulting in exposure of many individuals to HCHO on a daily basis. Although exposure levels may be low, chronic exposure may initiate respiratory disorders (Alexandersson and Hedenstierna, 1989) or possibly aggravate the effects of cigarette smoking, chronic bronchitis, or other respiratory illnesses (Krzyanowski et al., 1990). Early studies demonstrated HCHO injury to vital mucociliary clearance (CarThe authors thank Lisa P. Evans for technical assistance in manuscript preparation. This work was supported by grant R29ES04137 from NIEHS. Requests for reprints should be sent to Annette T. Hastie, Department of Medicine, Division of Pulmonary Medicine and Critical Care, Thomas Jefferson University, 1025 Walnut Street, Room 804, Philadelphia, PA 19107. 221 Journal of Toxicology and Environmental Health, 35:221-234, 1992 Copyright © 1992 by Hemisphere Publishing Corporation
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son et al., 1966; Cralley, 1942; Dalhamn, 1956; Falk et al., 1963). HCHO exposure causes lesions consisting of damaged epithelial cells corresponding to areas of ciliastasis and mucostasis (Morgan et al., 1986b). We and others have shown reversible ciliary dysfunction elicited by less extensive HCHO exposure (Hastie et al., 1990b; Kensler and Battista, 1966; Morgan et al., 1986a). Inhibition of zones of active epithelium is concentration and time dependent, and reduction of ciliary beat frequency is concentration dependent. Furthermore, we have found that treatment of tracheal epithelium with increasing HCHO concentrations (0, 16, 33, and 66 /xg/cm2 epithelial surface area) correspondingly reduces the yield of extractable cilia and significantly diminishes the specific ATPase activity of extracted ciliary axonemes, indicating loss of function. A recovery period results in a concentration-dependent return of ciliary function and in nearly normal amounts of extractable, functionally intact cilia (Hastie et al., 1990b). Therefore, HCHO-related injury appears to correlate with altered extraction of cilia from the epithelium. We have hypothesized that HCHO may elicit modifications of surface membrane components (Hastie et al., 1990b). Three glycoproteins with masses of 92, 98, and 105 kD are associated with ciliary membranes and are most readily accessible during labeling of the epithelium (Hastie et al., 1990a). Certain other proteins, accessible at the apical surface as evidenced by acquisition of biotin label, resist detergent solubilization of the lipid bilayer and remain associated with the internal ciliary axoneme (Hastie et al., 1990a). This investigation examines the HCHO-induced alterations of surface-accessible components in tracheal ciliated cells.
METHODS Biotinylation of Bovine Tracheal Epithelium
By permit from the U.S. Department of Agriculture, six bovine tracheae were obtained from a local slaughterhouse immediately after exsanguination of the animals. Equal halves of each trachea were rinsed twice with 0.9% sterile saline and incubated for 30 min at room temperature in 60 ml of 0.45 miW of the long-chain analog of N-hydroxysuccinimidobiotin (NHS-LC-biotin) in phosphate-buffered saline (PBS; 0.12 M NaCI, 2.7 miW KCI, 5 mJW potassium phosphate, pH 7.4) with mild agitation. Each tracheal half was rinsed briefly with two 50-ml aliquots of sterile saline. Tracheal Explant Treatment with HCHO
The two biotinylated tracheal halves were divided in half again directly after rinsing. Each quarter was submerged in PBS containing 0, 37.5, 75, or 150 /xg HCHO/ml for 30 min. This time period was selected
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on the basis of inhibition observed in rabbit tracheal rings. The extent of active ciliated epithelium was reduced 0, 5, 50, and 100% compared with that initially active at these concentrations after 30 min. If it is assumed that the total HCHO available reacts completely and evenly per unit surface area, HCHO concentrations of 37.5, 75, and 150 /ig/ml correspond to 16, 33, and 66 ng maximum per square centimeter of tracheal epithelial surface (Hastie et al., 1990b). This calculation, in fact, overestimates the actual concentration reacting with the epithelium by neglecting any HCHO remaining in solution at the end of the exposure period. A determination of HCHO concentration in the incubation solution immediately after exposure revealed that 77-81% remains after 30 min. Only 36-57% was found to remain after 60 min (Hastie et al., 1990b). For ease of comparison to our earlier work, the values are reported at the initial maximum HCHO concentration available per epithelial surface area: 0,16, 33, and 66 /ig/cm2. After incubation at 37°C for 30 min, the tracheal lengths were quickly rinsed with saline and the cilia were isolated. Isolation of Cilia from Epithelium and Fractionation into Ciliary Membranes and Axonemes
The method of isolation, described in detail by Hastie et al. (1990a), preserves the membrane surrounding the internal ciliary axoneme and can subsequently be separated into membrane + matrix and axoneme fractions. Briefly, each tracheal lumen is lightly brushed to disrupt the epithelium. The brush was rinsed with 40 ml of extraction buffer (50 mM NaCI, 20 mM Tris-HCI, pH 7.5, 1 mM EDTA, 7 mM 2-mercaptoethanol, and 10 mM CaCI2) containing 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxypropane-1-sulfonate detergent (CHAPS), which was poured into the tracheal lumen and vortexed for 30 s. The suspension of cellular debri and cilia from each tracheal quarter was retained and added to a 40-ml rinse of the lumen with extraction buffer without CHAPS. Each suspension was centrifuged at 2000 x g for 5 min and the supernate obtained was subjected to 12,000 x g for min. The pellet from the second centrifugation, containing the isolated cilia, was resuspended in 1 ml of resuspension buffer (RB; 50 mM potassium acetate, 20 mM Tris-HCI, pH 8.0, 4 mM MgSO 4 ,1 mM dithiothreitol, 0.5 mM EDTA) and repelleted at 12,000 x g for 1.5 min. The pellet was resuspended in 0.5 ml resuspension buffer with 0.1 mg/ml soybean trypsin inhibitor, and an aliquot of the cilia suspension was examined by light microscopy and recorded on videotape (Hastie et al., 1986). The cilia were pelleted at 12,000 x g for 1.5 min to fractionate into membrane and axoneme portions. The resulting pellet was resuspended in 0.5 ml of 0.5% Triton X-100 in resuspension buffer for 10 min at room temperature. The suspension was centrifuged for 1.5 min at
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12,000 x g . The supernate, containing the detergent soluble membrane + matrix fraction, was saved, and the pellet, containing the extracted ciliary axonemes, was resuspended in resuspension buffer with soybean trypsin inhibitor. Volumes of Triton membrane extraction and axoneme resuspension buffers were kept equal for each tracheal piece so that differences in yield could be noted.
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Treatment of Biotinylated Rabbit Tracheal Explants with HCHO
Excised rabbit trachea was biotinylated with NHS-LC-biotin at a concentration of 0.45 mM in phosphate-buffered saline at room temperature for 30 min. After incubation in biotin label, the trachea was rinsed with sterile saline, and tracheal ring explants of 1.2 mm thickness were cut. The tracheal explants were examined by light microscopy, and active ciliated epithelium in two separate zones per explant was video recorded. Four explants were then incubated in 0, 37.5, 75, and 150 ^g HCHO/ml for 1 h. Videorecordings of the same two zones in each explant were made every 10 min and assessed for decreased ciliary activity as described in detail (Hastie et al., 1987). The experiment was repeated with tracheae from three rabbits. Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE)
Equal volume aliquots of ciliary axoneme and membrane + matrix fractions were prepared for electrophoresis by diluting 1 : 1 with sample buffer and heating at 100°C for 2-3 min. Samples were electrophoresed in duplicate on SDS 4-10% polyacrylamide gradient gels using standard discontinuous, dissociating buffers (Maizel, 1971). One half of the gel slab was placed in Coomassie brilliant blue stain and the other half electrophoretically transferred onto Immobilon membrane (Millipore Corp., Bedford, Mass.) in 25 mJW Tris, 0.2 M glycine at 100 V for 1 h. Immobilon transfers were blocked, rinsed, and incubated with 1 : 500 dilution of streptavidin-biotin-horseradish peroxidase complex (Amersham Corp., Arlington Heights, III.) in Tris-buffered saline (18 mlW Tris, 0.5 M NaCI, pH 7.5) with 0.5% ovalbumin. The transfer was rinsed three times with Tris-buffered saline and developed for 10 min with 4chloro-1-naphthol. Both gels and transfers were scanned by a densitometer (Hoefer Scientific Instruments, San Francisco, Calif.) to quantitate the data. The baseline and gain were adjusted and held constant for each data set of ciliary axoneme or membrane + matrix samples from 0, 16, 33, and 66 Hg HCHO/cm2 exposure in each experiment. Densitometric analysis may become nonlinear at greater material loads. These may be underestimated and thus differences in comparison to lesser material loads may actually be greater than demonstrated. Molecular masses were estimated from migration of standard marker proteins.
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Protein Concentration, ATPase Activity, and HCHO Concentration Assays
Protein concentration was determined using the BioRad Protein MicroAssay (BioRad, Richmond, Calif.) based on the method of Bradford (1976) with ovalbumin as the standard protein. For ATPase assay, equal aliquots of axoneme suspensions were diluted into 200 /xl RB with 0.375 M NaCI plus 25jil of 20 mM ATP, 6 mM MgSO4, and were incubated for 30 min at 37°C. Axonemes were pelleted and the supernates were sampled in duplicate for inorganic phosphate (P|) by the Fiske and Subbarrow (1925) method. Specific ATPase activity was expressed as the ATPase activity per milligram protein. HCHO concentrations were determined by the chromotropic acid reagent method according to the NIOSH Manual of Analytical Methods (Eller, 1984). Statistical Analysis of Data
Linear regression analyses of dependent variables (protein and ATPase assays and densitometric results) versus HCHO concentration were performed using multivariate general linear hypothesis tests in the Systat 4.1 program. All trends reported as significant HCHO concentration-dependent changes had r values sufficient to reject that the regression line slope = 0. Tests for lack of fit in linear regression were conducted post hoc on those results reported to have a significant HCHO concentration-dependent effect. In all cases, it was determined that there was insufficient evidence to suggest that the linear regression model was inappropriate. RESULTS Decreased numbers of cilia were isolated from biotinylated tracheal epithelium exposed to increasing HCHO concentrations (Fig. 1). Correspondingly, axoneme fractions derived from Triton X-100 extraction of the isolated, biotin-labeled cilia also had significant HCHO concentration-dependent decreases in protein concentration, ATPase activity, and specific activity of ATPase (Table 1). These cilia were separated into a Triton X-100-soluble membrane + matrix fraction and a detergent-insoluble axoneme fraction. Surface accessible components identified by covalently linked biotin label segregate into both fractions (Hastie et al., 1990a). The membrane + matrix and axoneme subfractiohs of the cilia were examined by SDS-PAGE. Results from one of six experiments are shown in Figure 2. A decrease in the amount of axonemal proteins obtained, including tubulin (53-55 kD) and dynein (>300 kD), was observed; this corresponded with the decreased numbers of isolated cilia
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FIGURE 1. Light micrographs of isolated cilia from trachea quartered and exposed to (a) 0, (b) 16, (c) 33, and (cf) 66 ng HCHO/cm 2 for 30 min. Equal volumes of extraction and resuspension buffers were used. The yield of cilia decreased with increasing HCHO concentration. The micrographs are printed to the same magnification; the bar represents 10 f»m.
(see Fig. 1) and decreased total protein obtained after exposure to increasing HCHO concentrations (Table 1). The amount of protein in the membrane + matrix fractions remained approximately the same. Biotin-labeled components in ciliary membrane + matrix and axoneme fractions in the same samples displayed differing responses to increasing HCHO concentrations (Fig. 2). A substantial decline occurred in biotin-labeled material in the membrane + matrix fractions, TABLE 1. Axoneme Fraction Protein, ATPase, and Specific ATPase Content for Increasing HCHO Concentrations
HCHO (/ig/cm2)
Protein (mg/ml)
0 16 33 66
1.49 1.71 1.20 0.71
± ± ± ±
0.2 0.4 0.1 0.4
ATPase activity (nmol P|/min/ml)
Specific activity of ATPase (nmol P/min/mg protein)
514 589 414 160
349 356 341 231
± ± ± ±
160 12 93 87
± 110 ± 79 ±50 ± 29
Note. Values are means ± standard deviation. HCHO concentration-dependent decreases in yield of axoneme protein (r - -.73, p < .01), ATPase activity (r - - . 8 2 , p < .001), and specific activity of ATPase (r - -.58, p < .05) were statistically significant.
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D O 16 33 66 0 16 33 66
S
O 16 33 66 0 16
217K
33 66 S
200 K
116
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103
69
93
66
49
45 32
FIGURE 2. Representative SDS-PAGE analysis of ciliary membrane + matrix and axoneme fractions from tracheal quarters exposed to 0,16, 33, and 66 /*g HCHO/cm 2 in one of six experiments. This figure serves as a visual reference for the results presented: biotin-labeled components are detected in the transfer of axoneme (a) and membrane + matrix fractions (b), the proteins are shown in the stained gel of those same axoneme (c) and membrane + matrix fractions (d), migration of tubulin (T) and dynein (D) proteins are marked, biotin-labeled axoneme components at 200 and 150 kD are indicated by arrowheads on the left of A, biotin-labeled membrane components at 92, 98, and 105 kD are indicated by open dots at the right of B, and the molecular weights of the standards (s) are given in kilodaltons on the right of the transfer and the gel.
particularly in the major surface-accessible components identified previously at 92, 98, and 105 kD (Hastie et al., 1990a), corresponding to increasing HCHO exposure. These components stain lightly with Coomassie brilliant blue, indicating that they were present in low amounts. Two other biotin-labeled components in the membrane + matrix fraction, one at 126 and a doublet at 76 kD, were found previously to bind streptavidin without prior biotin labeling and to remain in the supernate instead of sedimenting with reconstituted membrane vesicles (Hastie et al., 1990a). Thus these two were presumably not membrane constituents, despite their seeming acquisition of biotin label. Although certain biotin-labeled components in the ciliary axonemes (e.g., those at 180 and >300 kD) were diminished substantially at greater HCHO concentrations (see Fig. 2a), there was little decline in the total amount of biotin-labeled material, in contrast to results in the membrane + matrix fraction. In fact, some surface-accessible, biotinlabeled proteins remaining with the axoneme (e.g., those at 150 and 200
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kD) were increased significantly in a HCHO concentration-dependent trend (Table 2). Therefore, the association of these surface-accessible proteins with the axoneme or their resistance to detergent solubilization was enhanced by exposure to HCHO. Densitometric analyses of CBB stained total protein profiles in ciliary axoneme and membrane + matrix fractions from the six experiments are presented in Figure 3. There was a significant decline in total axoneme protein with increasing HCHO concentration (r = -.66, p = .001), but not in membrane + matrix protein. Specific ciliary axoneme structural proteins, tubulin and dynein, were also significantly reduced with increasing HCHO concentrations (r = -.75, p < .001, and r = -.45, p = .03, respectively). The largest standard deviation occurs in the control fraction (0 HCHO concentration), indicating the variability between the different trachea, most likely in the extent of ciliated cells present. Densitometric analyses of biotin-labeled material in ciliary axoneme and membrane + matrix fractions from the six experiments are shown in Figure 3. There was a significant decrease in biotin-labeled material in the membrane + matrix fraction (r — —.47, p = .019). Biotin-labeled material in the axoneme fraction did not decrease significantly. The proportion of biotin-labeled material with respect to total protein in the axoneme fraction increased significantly with increasing HCHO concentration (r = — .47, p = .03). The proportion of biotin-labeled material to total protein in the membrane + matrix fraction declined, but not significantly (r = -.36, p = .08) (Fig. 4). DISCUSSION These results provide additional new information at a molecular level concerning the mechanism of HCHO-induced ciliary dysfunction. TABLE 2. Average Percent of Control Level for Specific Biotin-Labeled Proteins and Total BiotinLabeled Material in Axoneme Fractions at Increasing HCHO Concentrations as Determined from SDS-PAGE HCHO (/ig/cm2)
200-kD Biotin protein
150-kD Biotin protein
Total Biotin protein
0 16 33 66
100% 120 ± 38% 172 ± 46% 188 ± 92%
100% 105 ± 29%
100%
141 ±20% 144 ± 59%
95 ± 13% 104 ± 18% 98 ± 10%
Afofe. Densitometric units for specific proteins or total were expressed as a percent of control (0 /jg/cm2 HCHO) level. Means ± standard deviations were calculated from four experiments. There were significant HCHO concentration-dependent increases in biotin-labeled proteins at 200 and 150 kD (r — .59, p < .02, and r - .52, p < .04, respectively), but no significant change in the overall total of biotin-labeled material in the axoneme fractions.
Axonemes -
CBB
Axonemes o
-
Biotin
50 40
uni
CO
— L
30
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o
'B
20
E o 10 "3> 0
10
20
30
40
50
60
70
2
HCOH concentration (ug/cm )
Membranes
10
-
20
0
0
10
Membranes
40
HCOH concentration
50
60 2
(ug/cm )
20
30
40
HCOH concentration
CBB
30
V •o
70
10
-
20
50
60
70
(ug/cm*)
Biotin
30
40
HCOH concentration
50
60 2
(ug/cm )
FIGURE 3. Densitometric analyses of Coomassie brilliant blue (CBB) stained total protein profiles in ciliary axonemes and membrane + matrix samples are presented as means and standard deviations from six experiments in the graphs on the left. There was a significant decline in the mean total axonemal protein (r - -.66, p - .001) but not in the mean total membrane + matrix protein. Densitometric analyses of biotin-labeled material in ciliary axoneme and membrane + matrix samples are presented as means and standard deviations from six experiments in the graphs on the right. There was a significant decrease in biotin-labeled material in the membrane + matrix fraction (r - -A7, p - .019) but not in the axoneme fraction.
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Axonemes
ID O
10
20
30
40
SO
60
70
HCOH concentration (ug/cm2) Membranes co 4
0
10
20
30
40
50
60
70
HCOH concentration (ug/cm1) FIGURE 4. Ratios of biotin-labeled material to total protein in ciliary axoneme and membrane + matrix samples are presented as means and standard deviations from six experiments. The proportion of biotin-labeled material increases significantly in the axoneme fractions (r — —.47, p — .03). Biotin-labeled material declined in the membrane fractions but not significantly.
The yield of cilia with surrounding membranes extractable from HCHOexposed epithelium decreased in numbers with increasing HCHO concentration. The axoneme subfraction from these isolated cilia yielded decreasing protein concentration, ATPase activity, and specific ATPase activity with increasing HCHO concentration. The measured decrease in protein of the axoneme fraction was confirmed by densitometric analysis of total protein and axoneme specific proteins, tubulin and dynein, in samples on SDS-PAGE. These results from biotin-labeled tracheal epithelium are in complete agreement with previous work on nonlabeled epithelium (Hastie et al., 1990b). Because HCHO exposure impaired the extraction of ciliary axonemes by techniques of membrane perturbation, CaCI2 influx and
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shear, we hypothesized that alterations elicited by HCHO might involve surface membrane components (Hastie et al., 1990b). In previous studies, the membranes were solubilized during the axoneme isolation procedure and thus could not be examined (Hastie et al., 1986). However, with modification of the isolation buffer and method, the ciliary membranes were retained as a fraction that could be investigated further (Hastie et al., 1990a). Biotin was conjugated covalently to reactive sites on the intact epithelium prior to HCHO exposure to identify surface-accessible components. There are some membrane constituents that are inaccessible or unreactive for attachment of label (Hastie et al., 1990a). However, biotin labeling of membranes in epithelial cells has revealed greater numbers of constituents than metabolic labeling methods (Lisanti et al., 1989). The modified method for extraction of cilia, although retaining ciliary membranes, introduces some additional contaminants including the internal, soluble "matrix" proteins of the cilia (Gibbons, 1965), which are released into the membrane fraction by detergent solubilization of the lipid bilayer and do not associate with reconstituted membranes (Hastie et al., 1990a). Thus biotin labeling was considered necessary to distinguish external components of ciliary membranes. Labeling with biotin did not modify subsequent epithelial response to HCHO as evidenced by similar concentration- and time-dependent inhibition of functionally active, biotinylated rabbit tracheal epithelium compared with tracheal explants that were not biotin labeled. Moreover, equivalent decreased yields of extractable cilia from biotin-labeled and nonlabeled epithelium indicate that the HCHO concentrationdependent alterations in the epithelium were not influenced by the presence of biotin label, nor by the modified procedure for isolation of cilia. The HCHO concentration-dependent decline in extractable cilia predictably resulted in reduced membrane material as well. This was demonstrated by significantly decreased biotin-labeled components identifying surface-accessible material in the detergent-soluble membrane + matrix fraction. The total protein content in the membrane + matrix fractions analyzed by SDS-PAGE remained unchanged. However, the majority of these proteins are not biotinylated, indicating that they were probably not surface-accessible components. In addition, many of these same proteins remain in the supernate upon sedimentation of reconstituted membrane vesicles (Hastie et al., 1990a), which is further evidence that these proteins were not membrane constituents. As stated earlier, nonlabeled proteins in the membrane + matrix fraction may consist of the soluble, internal matrix proteins of cilia. Although decreased numbers of extracted cilia would predictably yield less of these matrix proteins as well, decreased permeability of the ciliary membrane would ensure,
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conversely, that more of these proteins remained within the cilia instead of being lost during the cilia isolation procedure. The biotinylated components in the axoneme fractions, by virtue of possessing this covalently linked label, are accessible to the external environment. The majority of these is distinctly different from biotinylated components in the membrane + matrix fractions and resists detergent solubilization, indicating a close association with the internal ciliary axoneme (Hastie et al., 1990a). The total amount of biotin-labeled material in the axoneme fraction remained nearly the same with increasing HCHO concentration. Thus, considered as a proportion of the total protein in the axoneme fraction, biotinylated material increased significantly with increasing HCHO concentration. This trend was supported by significant, concentration-dependent increases in specific biotin-labeled proteins in the axoneme fraction, although other biotinlabeled proteins in this fraction decreased. Thus not all surfaceaccessible components were affected by HCHO in the same way or to the same extent. Casanova-Schmitz et al. (1983) observed decreased extractability of DNA from protein in rat nasal mucosa with increasing HCHO exposure. The increased resistance to normal extraction procedures for cilia from the epithelium and for certain surface-accessible components from association with the axoneme after HCHO exposure may be related phenomena. The pattern of protein migration did not noticeably change with increased HCHO exposure, indicating that molecular weights of the components remained the same. Although the increase of biotinlabeled components in the axoneme fraction might possibly derive from cross-linking of smaller biotin-labeled components, it is more likely that these would appear as new bands, instead of migrating to precisely the same positions as other components already present. Thus covalent linkages between proteins apparently were not formed by HCHO exposure. However, internal methylene bridge formation between amino groups within a protein molecule rather than crosslinking between adjacent molecules (French and Edsall, 1945) may make the epithelial surface components less susceptible to detergent solubilization. In conclusion, we have demonstrated (1) reduced extraction of surface accessible membrane components; (2) increased retention of internal soluble proteins within the cilia, subsequently released into the membrane + matrix fraction; and (3) increased retention of surface accessible components with internal axonemes. Together, these points confirm that components accessible at the epithelial surface were altered to varying degrees by exposure to HCHO, possibly through internal molecular stabilization. Such alterations, if not reversed, may result in other secondary responses leading to loss of cilia and to cell injury and death, which have been observed at higher HCHO concentrations
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or longer exposure periods (Bhalla et al., 1991; Monteiro-Riviere and Popp, 1986; Monticello et al., 1989; Zwart et al., 1988).
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REFERENCES Alexandersson, R., and Hedenstierna, G. 1989. Pulmonary function in wood workers exposed to formaldehyde: A prospective study. Arch. Environ. Health 44:5-11. Bhalla, D. K., Mahavni, V., Nguyen, T., and McClure, T. 1991. Effects of acute exposure to formaldehyde on surface morphology of nasal epithelia in rats. J. Toxicol. Environ. Health 33:171-188. Bradford, M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. Carson, S., Goldhamer, R., and Carpenter, R. 1966. Mucus transport in the respiratory tract. Am. Rev. Respir. Dis. 93:86-92. Casanova-Schmitz, M., and Heck, H. D'A. 1983. Effects of formaldehyde exposure on the extractability of DNA from proteins in the rat nasal mucosa. Toxicol. Appl. Pharmacol. 70:121-132. Cralley, L. V. 1942. The effect of irritant gases upon the rate of ciliary activity. J. Ind. Hyg. Toxicol. 24:193-198. Dalhamn, T. 1956. Mucous flow and ciliary activity in the trachea of healthy rats and rats exposed to respiratory irritant gases (SO2, N 3 H, HCHO). Acta Physiol. Scand. 36(Suppl. 123):1-161. Eller, P. M., ed. 1984. NIOSH Manual of Analytical Methods, 3d ed., pp. 3500-1-3500-4. Washington, D.C.: Government Printing Office. Publication number 84-100. Falk, H. L., Kotin, P., and Rowlette, W. 1963. The response of mucus-secreting epithelium and mucus to irritants. Ann. N.Y. Acad. Sci. 106:583-608. Fiske, C. H., and Subbarrow, Y. 1925. The colorimetric determination of phosphorus, f. Biol. Chem. 66:375-400. French, D., and Edsall, J. T. 1945. The reactions of formaldehyde with amino acids and proteins. Adv. Protein Chem. 2:277-335. Gibbons, I. R. 1965. Chemical dissection of cilia. Arch. Biol. (Liege) 76:317-352. Hastie, A. T., Dicker, D. T., Hingley, S. I , Kueppers, F., Higgins, M. L., and Weinbaum, G. 1986. Isolation of cilia from porcine tracheal epithelium and extraction of dynein arms. Cell Motil. Cytoskel. 6:25-34. Hastie, A. T., Loegering, D. A., Gleich, G. J., and Kueppers, F. 1987. The effect of purified human eosinophil major basic protein on mammalian ciliary activity. Am. Rev. Respir. Dis. 135:848853. Hastie, A. T., Krantz, M. J., and Colizzo, F. P. 1990a. Identification of surface components of mammalian respiratory tract cilia. Cell Motil. Cytoskel. 17:317-328. Hastie, A. T., Patrick, H., and Fish, J. E. 1990b. Inhibition and recovery of mammalian respiratory ciliary function after formaldehyde exposure. Toxicol. Appl. Pharmacol. 102:282-291. Kensler, C. J., and Battista, S. P. 1966. Chemical and physical factors affecting mammalian ciliary activity. Am. Rev. Respir. Dis. 93:93-102. Krzyanowski, M., Quackenboss, J. J., and Lebowitz, M. D. 1990. Chronic respiratory effect of indoor formaldehyde exposure. Environ. Res. 52:117-125. Lisanti, M., Le Bivic, A., Sargiacomo, M., and Rodriguez-Boulan, E. 1989. Steady-state distribution and biogenesis of endogenous Madin-Darby canine kidney glycoproteins: Evidence for intracellular sorting and polarized cell surface delivery. J. Cell Biol. 109:2117-2127. Maizel, J. V., Jr. 1971. Polyacrylamide gel electrophoresis of viral proteins. Methods Virol. 5:179-246. Monterio-Riviere, N. A., and Popp, J. A. 1986. Ultrastructural evaluation of acute nasal toxicity in the rat respiratory epithelium in response to formaldehyde gas. Fundam. Appl. Toxicol. 6:251262. Monticello, T. M., Morgan, K. T., Everitt, J. I., and Popp, J. A. 1989. Effects of formaldehyde gas on the respiratory tract of rhesus monkeys. Am. J. Pathol. 134:515-527. Morgan, K. T., Gross, E. A., and Patterson, D. L. 1986a. Distribution, progression, and recovery of
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acute formaldehyde-induced inhibition of nasal mucociliary function in F-344 rats. Toxicol. Appl. Pharmacol. 86:448-456. Morgan, K. T., Patterson, D. L., and Gross, E. A. 1986b. Responses of the nasal mucociliary apparatus of F-344 rats to formaldehyde gas. Toxicol. Appl. Pharmacol. 82:1-13. Zwart, A., Woutersen, R. A., Wilmer, J. W. C. M., Spit, B. J., and Feron, V. J. 1988. Cytotoxic and adaptive effects in rat nasal epithelium after 3 day and 13 week exposure to low concentrations of formaldehyde vapour. Toxicology 51:87-99.
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Received August 10, 1991 Accepted November 15, 1991
ISSN: 0098-4108 (Print) (Online) Journal homepage: http://www.tandfonline.com/loi/uteh19
Ciliated respiratory epithelial surface changes after formaldehyde exposure Frank Colizzo , Mori J. Krantz , James E. Fish & Annette T. Hastie To cite this article: Frank Colizzo , Mori J. Krantz , James E. Fish & Annette T. Hastie (1992) Ciliated respiratory epithelial surface changes after formaldehyde exposure, Journal of Toxicology and Environmental Health, 35:4, 221-234, DOI: 10.1080/15287399209531613 To link to this article: http://dx.doi.org/10.1080/15287399209531613
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Date: 07 November 2015, At: 22:54
CILIATED RESPIRATORY EPITHELIAL SURFACE CHANGES AFTER FORMALDEHYDE EXPOSURE Frank Colizzo, Mori J. Krantz, James E. Fish, Annette T. Hastie
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Department of Medicine, Thomas Jefferson University, Philadelphia, Pennsylvania
The investigation sought to identify alterations of specific ciliated epithelial surface components after exposure to formaldehyde (HCHO) levels that decrease respiratory ciliary function. Bovine tracheae were reacted with an analog of N-hydroxysuccinimidobiotin to label epithelial surface-accessible components before exposure to HCHO. The tracheae were then exposed to 0, 16, 33, and 66 ng HCHO/cm2 epithelial surface for 30 min. Cilia were isolated from the epithelium, separated into membrane and internal axonemal portions, analyzed on sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PACE), and either stained to detect proteins or transblotted to detect biotin-labeled components. Densitometric analysis of axoneme proteins showed a decrease in the total amount extracted with increased HCHO concentration, including axoneme-specific proteins, dynein, and tubulin. However, biotinylated proteins in the axoneme fractions proportionately increased. Membrane fractions showed little change in protein with increasing HCHO concentration. The majority of these is not biotin-labeled and thus not surface-accessible components. Biotinylated material in the membrane fractions showed a significant decrease with increased HCHO concentration, particularly of bands at 92, 98, and 105 kD. These data suggest that increasing HCHO exposure reduces both extractable ciliary axonemes and detergent-soluble surface components, possibly by stabilizing respiratory epithelial membranes. This process apparently strengthens association of certain surface components with the internal axoneme, thereby reducing subsequent solubilization in detergent.
INTRODUCTION Formaldehyde (HCHO) occurs widely in industrial and domestic environments, resulting in exposure of many individuals to HCHO on a daily basis. Although exposure levels may be low, chronic exposure may initiate respiratory disorders (Alexandersson and Hedenstierna, 1989) or possibly aggravate the effects of cigarette smoking, chronic bronchitis, or other respiratory illnesses (Krzyanowski et al., 1990). Early studies demonstrated HCHO injury to vital mucociliary clearance (CarThe authors thank Lisa P. Evans for technical assistance in manuscript preparation. This work was supported by grant R29ES04137 from NIEHS. Requests for reprints should be sent to Annette T. Hastie, Department of Medicine, Division of Pulmonary Medicine and Critical Care, Thomas Jefferson University, 1025 Walnut Street, Room 804, Philadelphia, PA 19107. 221 Journal of Toxicology and Environmental Health, 35:221-234, 1992 Copyright © 1992 by Hemisphere Publishing Corporation
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son et al., 1966; Cralley, 1942; Dalhamn, 1956; Falk et al., 1963). HCHO exposure causes lesions consisting of damaged epithelial cells corresponding to areas of ciliastasis and mucostasis (Morgan et al., 1986b). We and others have shown reversible ciliary dysfunction elicited by less extensive HCHO exposure (Hastie et al., 1990b; Kensler and Battista, 1966; Morgan et al., 1986a). Inhibition of zones of active epithelium is concentration and time dependent, and reduction of ciliary beat frequency is concentration dependent. Furthermore, we have found that treatment of tracheal epithelium with increasing HCHO concentrations (0, 16, 33, and 66 /xg/cm2 epithelial surface area) correspondingly reduces the yield of extractable cilia and significantly diminishes the specific ATPase activity of extracted ciliary axonemes, indicating loss of function. A recovery period results in a concentration-dependent return of ciliary function and in nearly normal amounts of extractable, functionally intact cilia (Hastie et al., 1990b). Therefore, HCHO-related injury appears to correlate with altered extraction of cilia from the epithelium. We have hypothesized that HCHO may elicit modifications of surface membrane components (Hastie et al., 1990b). Three glycoproteins with masses of 92, 98, and 105 kD are associated with ciliary membranes and are most readily accessible during labeling of the epithelium (Hastie et al., 1990a). Certain other proteins, accessible at the apical surface as evidenced by acquisition of biotin label, resist detergent solubilization of the lipid bilayer and remain associated with the internal ciliary axoneme (Hastie et al., 1990a). This investigation examines the HCHO-induced alterations of surface-accessible components in tracheal ciliated cells.
METHODS Biotinylation of Bovine Tracheal Epithelium
By permit from the U.S. Department of Agriculture, six bovine tracheae were obtained from a local slaughterhouse immediately after exsanguination of the animals. Equal halves of each trachea were rinsed twice with 0.9% sterile saline and incubated for 30 min at room temperature in 60 ml of 0.45 miW of the long-chain analog of N-hydroxysuccinimidobiotin (NHS-LC-biotin) in phosphate-buffered saline (PBS; 0.12 M NaCI, 2.7 miW KCI, 5 mJW potassium phosphate, pH 7.4) with mild agitation. Each tracheal half was rinsed briefly with two 50-ml aliquots of sterile saline. Tracheal Explant Treatment with HCHO
The two biotinylated tracheal halves were divided in half again directly after rinsing. Each quarter was submerged in PBS containing 0, 37.5, 75, or 150 /xg HCHO/ml for 30 min. This time period was selected
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on the basis of inhibition observed in rabbit tracheal rings. The extent of active ciliated epithelium was reduced 0, 5, 50, and 100% compared with that initially active at these concentrations after 30 min. If it is assumed that the total HCHO available reacts completely and evenly per unit surface area, HCHO concentrations of 37.5, 75, and 150 /ig/ml correspond to 16, 33, and 66 ng maximum per square centimeter of tracheal epithelial surface (Hastie et al., 1990b). This calculation, in fact, overestimates the actual concentration reacting with the epithelium by neglecting any HCHO remaining in solution at the end of the exposure period. A determination of HCHO concentration in the incubation solution immediately after exposure revealed that 77-81% remains after 30 min. Only 36-57% was found to remain after 60 min (Hastie et al., 1990b). For ease of comparison to our earlier work, the values are reported at the initial maximum HCHO concentration available per epithelial surface area: 0,16, 33, and 66 /ig/cm2. After incubation at 37°C for 30 min, the tracheal lengths were quickly rinsed with saline and the cilia were isolated. Isolation of Cilia from Epithelium and Fractionation into Ciliary Membranes and Axonemes
The method of isolation, described in detail by Hastie et al. (1990a), preserves the membrane surrounding the internal ciliary axoneme and can subsequently be separated into membrane + matrix and axoneme fractions. Briefly, each tracheal lumen is lightly brushed to disrupt the epithelium. The brush was rinsed with 40 ml of extraction buffer (50 mM NaCI, 20 mM Tris-HCI, pH 7.5, 1 mM EDTA, 7 mM 2-mercaptoethanol, and 10 mM CaCI2) containing 0.1% 3-[(3-cholamidopropyl)dimethylammonio]-2-hydroxypropane-1-sulfonate detergent (CHAPS), which was poured into the tracheal lumen and vortexed for 30 s. The suspension of cellular debri and cilia from each tracheal quarter was retained and added to a 40-ml rinse of the lumen with extraction buffer without CHAPS. Each suspension was centrifuged at 2000 x g for 5 min and the supernate obtained was subjected to 12,000 x g for min. The pellet from the second centrifugation, containing the isolated cilia, was resuspended in 1 ml of resuspension buffer (RB; 50 mM potassium acetate, 20 mM Tris-HCI, pH 8.0, 4 mM MgSO 4 ,1 mM dithiothreitol, 0.5 mM EDTA) and repelleted at 12,000 x g for 1.5 min. The pellet was resuspended in 0.5 ml resuspension buffer with 0.1 mg/ml soybean trypsin inhibitor, and an aliquot of the cilia suspension was examined by light microscopy and recorded on videotape (Hastie et al., 1986). The cilia were pelleted at 12,000 x g for 1.5 min to fractionate into membrane and axoneme portions. The resulting pellet was resuspended in 0.5 ml of 0.5% Triton X-100 in resuspension buffer for 10 min at room temperature. The suspension was centrifuged for 1.5 min at
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12,000 x g . The supernate, containing the detergent soluble membrane + matrix fraction, was saved, and the pellet, containing the extracted ciliary axonemes, was resuspended in resuspension buffer with soybean trypsin inhibitor. Volumes of Triton membrane extraction and axoneme resuspension buffers were kept equal for each tracheal piece so that differences in yield could be noted.
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Treatment of Biotinylated Rabbit Tracheal Explants with HCHO
Excised rabbit trachea was biotinylated with NHS-LC-biotin at a concentration of 0.45 mM in phosphate-buffered saline at room temperature for 30 min. After incubation in biotin label, the trachea was rinsed with sterile saline, and tracheal ring explants of 1.2 mm thickness were cut. The tracheal explants were examined by light microscopy, and active ciliated epithelium in two separate zones per explant was video recorded. Four explants were then incubated in 0, 37.5, 75, and 150 ^g HCHO/ml for 1 h. Videorecordings of the same two zones in each explant were made every 10 min and assessed for decreased ciliary activity as described in detail (Hastie et al., 1987). The experiment was repeated with tracheae from three rabbits. Sodium Dodecyl Sulfate Polyacrylamide Gel Electrophoresis (SDS-PAGE)
Equal volume aliquots of ciliary axoneme and membrane + matrix fractions were prepared for electrophoresis by diluting 1 : 1 with sample buffer and heating at 100°C for 2-3 min. Samples were electrophoresed in duplicate on SDS 4-10% polyacrylamide gradient gels using standard discontinuous, dissociating buffers (Maizel, 1971). One half of the gel slab was placed in Coomassie brilliant blue stain and the other half electrophoretically transferred onto Immobilon membrane (Millipore Corp., Bedford, Mass.) in 25 mJW Tris, 0.2 M glycine at 100 V for 1 h. Immobilon transfers were blocked, rinsed, and incubated with 1 : 500 dilution of streptavidin-biotin-horseradish peroxidase complex (Amersham Corp., Arlington Heights, III.) in Tris-buffered saline (18 mlW Tris, 0.5 M NaCI, pH 7.5) with 0.5% ovalbumin. The transfer was rinsed three times with Tris-buffered saline and developed for 10 min with 4chloro-1-naphthol. Both gels and transfers were scanned by a densitometer (Hoefer Scientific Instruments, San Francisco, Calif.) to quantitate the data. The baseline and gain were adjusted and held constant for each data set of ciliary axoneme or membrane + matrix samples from 0, 16, 33, and 66 Hg HCHO/cm2 exposure in each experiment. Densitometric analysis may become nonlinear at greater material loads. These may be underestimated and thus differences in comparison to lesser material loads may actually be greater than demonstrated. Molecular masses were estimated from migration of standard marker proteins.
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Protein Concentration, ATPase Activity, and HCHO Concentration Assays
Protein concentration was determined using the BioRad Protein MicroAssay (BioRad, Richmond, Calif.) based on the method of Bradford (1976) with ovalbumin as the standard protein. For ATPase assay, equal aliquots of axoneme suspensions were diluted into 200 /xl RB with 0.375 M NaCI plus 25jil of 20 mM ATP, 6 mM MgSO4, and were incubated for 30 min at 37°C. Axonemes were pelleted and the supernates were sampled in duplicate for inorganic phosphate (P|) by the Fiske and Subbarrow (1925) method. Specific ATPase activity was expressed as the ATPase activity per milligram protein. HCHO concentrations were determined by the chromotropic acid reagent method according to the NIOSH Manual of Analytical Methods (Eller, 1984). Statistical Analysis of Data
Linear regression analyses of dependent variables (protein and ATPase assays and densitometric results) versus HCHO concentration were performed using multivariate general linear hypothesis tests in the Systat 4.1 program. All trends reported as significant HCHO concentration-dependent changes had r values sufficient to reject that the regression line slope = 0. Tests for lack of fit in linear regression were conducted post hoc on those results reported to have a significant HCHO concentration-dependent effect. In all cases, it was determined that there was insufficient evidence to suggest that the linear regression model was inappropriate. RESULTS Decreased numbers of cilia were isolated from biotinylated tracheal epithelium exposed to increasing HCHO concentrations (Fig. 1). Correspondingly, axoneme fractions derived from Triton X-100 extraction of the isolated, biotin-labeled cilia also had significant HCHO concentration-dependent decreases in protein concentration, ATPase activity, and specific activity of ATPase (Table 1). These cilia were separated into a Triton X-100-soluble membrane + matrix fraction and a detergent-insoluble axoneme fraction. Surface accessible components identified by covalently linked biotin label segregate into both fractions (Hastie et al., 1990a). The membrane + matrix and axoneme subfractiohs of the cilia were examined by SDS-PAGE. Results from one of six experiments are shown in Figure 2. A decrease in the amount of axonemal proteins obtained, including tubulin (53-55 kD) and dynein (>300 kD), was observed; this corresponded with the decreased numbers of isolated cilia
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FIGURE 1. Light micrographs of isolated cilia from trachea quartered and exposed to (a) 0, (b) 16, (c) 33, and (cf) 66 ng HCHO/cm 2 for 30 min. Equal volumes of extraction and resuspension buffers were used. The yield of cilia decreased with increasing HCHO concentration. The micrographs are printed to the same magnification; the bar represents 10 f»m.
(see Fig. 1) and decreased total protein obtained after exposure to increasing HCHO concentrations (Table 1). The amount of protein in the membrane + matrix fractions remained approximately the same. Biotin-labeled components in ciliary membrane + matrix and axoneme fractions in the same samples displayed differing responses to increasing HCHO concentrations (Fig. 2). A substantial decline occurred in biotin-labeled material in the membrane + matrix fractions, TABLE 1. Axoneme Fraction Protein, ATPase, and Specific ATPase Content for Increasing HCHO Concentrations
HCHO (/ig/cm2)
Protein (mg/ml)
0 16 33 66
1.49 1.71 1.20 0.71
± ± ± ±
0.2 0.4 0.1 0.4
ATPase activity (nmol P|/min/ml)
Specific activity of ATPase (nmol P/min/mg protein)
514 589 414 160
349 356 341 231
± ± ± ±
160 12 93 87
± 110 ± 79 ±50 ± 29
Note. Values are means ± standard deviation. HCHO concentration-dependent decreases in yield of axoneme protein (r - -.73, p < .01), ATPase activity (r - - . 8 2 , p < .001), and specific activity of ATPase (r - -.58, p < .05) were statistically significant.
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D O 16 33 66 0 16 33 66
S
O 16 33 66 0 16
217K
33 66 S
200 K
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69
93
66
49
45 32
FIGURE 2. Representative SDS-PAGE analysis of ciliary membrane + matrix and axoneme fractions from tracheal quarters exposed to 0,16, 33, and 66 /*g HCHO/cm 2 in one of six experiments. This figure serves as a visual reference for the results presented: biotin-labeled components are detected in the transfer of axoneme (a) and membrane + matrix fractions (b), the proteins are shown in the stained gel of those same axoneme (c) and membrane + matrix fractions (d), migration of tubulin (T) and dynein (D) proteins are marked, biotin-labeled axoneme components at 200 and 150 kD are indicated by arrowheads on the left of A, biotin-labeled membrane components at 92, 98, and 105 kD are indicated by open dots at the right of B, and the molecular weights of the standards (s) are given in kilodaltons on the right of the transfer and the gel.
particularly in the major surface-accessible components identified previously at 92, 98, and 105 kD (Hastie et al., 1990a), corresponding to increasing HCHO exposure. These components stain lightly with Coomassie brilliant blue, indicating that they were present in low amounts. Two other biotin-labeled components in the membrane + matrix fraction, one at 126 and a doublet at 76 kD, were found previously to bind streptavidin without prior biotin labeling and to remain in the supernate instead of sedimenting with reconstituted membrane vesicles (Hastie et al., 1990a). Thus these two were presumably not membrane constituents, despite their seeming acquisition of biotin label. Although certain biotin-labeled components in the ciliary axonemes (e.g., those at 180 and >300 kD) were diminished substantially at greater HCHO concentrations (see Fig. 2a), there was little decline in the total amount of biotin-labeled material, in contrast to results in the membrane + matrix fraction. In fact, some surface-accessible, biotinlabeled proteins remaining with the axoneme (e.g., those at 150 and 200
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kD) were increased significantly in a HCHO concentration-dependent trend (Table 2). Therefore, the association of these surface-accessible proteins with the axoneme or their resistance to detergent solubilization was enhanced by exposure to HCHO. Densitometric analyses of CBB stained total protein profiles in ciliary axoneme and membrane + matrix fractions from the six experiments are presented in Figure 3. There was a significant decline in total axoneme protein with increasing HCHO concentration (r = -.66, p = .001), but not in membrane + matrix protein. Specific ciliary axoneme structural proteins, tubulin and dynein, were also significantly reduced with increasing HCHO concentrations (r = -.75, p < .001, and r = -.45, p = .03, respectively). The largest standard deviation occurs in the control fraction (0 HCHO concentration), indicating the variability between the different trachea, most likely in the extent of ciliated cells present. Densitometric analyses of biotin-labeled material in ciliary axoneme and membrane + matrix fractions from the six experiments are shown in Figure 3. There was a significant decrease in biotin-labeled material in the membrane + matrix fraction (r — —.47, p = .019). Biotin-labeled material in the axoneme fraction did not decrease significantly. The proportion of biotin-labeled material with respect to total protein in the axoneme fraction increased significantly with increasing HCHO concentration (r = — .47, p = .03). The proportion of biotin-labeled material to total protein in the membrane + matrix fraction declined, but not significantly (r = -.36, p = .08) (Fig. 4). DISCUSSION These results provide additional new information at a molecular level concerning the mechanism of HCHO-induced ciliary dysfunction. TABLE 2. Average Percent of Control Level for Specific Biotin-Labeled Proteins and Total BiotinLabeled Material in Axoneme Fractions at Increasing HCHO Concentrations as Determined from SDS-PAGE HCHO (/ig/cm2)
200-kD Biotin protein
150-kD Biotin protein
Total Biotin protein
0 16 33 66
100% 120 ± 38% 172 ± 46% 188 ± 92%
100% 105 ± 29%
100%
141 ±20% 144 ± 59%
95 ± 13% 104 ± 18% 98 ± 10%
Afofe. Densitometric units for specific proteins or total were expressed as a percent of control (0 /jg/cm2 HCHO) level. Means ± standard deviations were calculated from four experiments. There were significant HCHO concentration-dependent increases in biotin-labeled proteins at 200 and 150 kD (r — .59, p < .02, and r - .52, p < .04, respectively), but no significant change in the overall total of biotin-labeled material in the axoneme fractions.
Axonemes -
CBB
Axonemes o
-
Biotin
50 40
uni
CO
— L
30
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o
'B
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E o 10 "3> 0
10
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HCOH concentration (ug/cm )
Membranes
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HCOH concentration
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HCOH concentration
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V •o
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-
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(ug/cm*)
Biotin
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(ug/cm )
FIGURE 3. Densitometric analyses of Coomassie brilliant blue (CBB) stained total protein profiles in ciliary axonemes and membrane + matrix samples are presented as means and standard deviations from six experiments in the graphs on the left. There was a significant decline in the mean total axonemal protein (r - -.66, p - .001) but not in the mean total membrane + matrix protein. Densitometric analyses of biotin-labeled material in ciliary axoneme and membrane + matrix samples are presented as means and standard deviations from six experiments in the graphs on the right. There was a significant decrease in biotin-labeled material in the membrane + matrix fraction (r - -A7, p - .019) but not in the axoneme fraction.
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Axonemes
ID O
10
20
30
40
SO
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HCOH concentration (ug/cm2) Membranes co 4
0
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HCOH concentration (ug/cm1) FIGURE 4. Ratios of biotin-labeled material to total protein in ciliary axoneme and membrane + matrix samples are presented as means and standard deviations from six experiments. The proportion of biotin-labeled material increases significantly in the axoneme fractions (r — —.47, p — .03). Biotin-labeled material declined in the membrane fractions but not significantly.
The yield of cilia with surrounding membranes extractable from HCHOexposed epithelium decreased in numbers with increasing HCHO concentration. The axoneme subfraction from these isolated cilia yielded decreasing protein concentration, ATPase activity, and specific ATPase activity with increasing HCHO concentration. The measured decrease in protein of the axoneme fraction was confirmed by densitometric analysis of total protein and axoneme specific proteins, tubulin and dynein, in samples on SDS-PAGE. These results from biotin-labeled tracheal epithelium are in complete agreement with previous work on nonlabeled epithelium (Hastie et al., 1990b). Because HCHO exposure impaired the extraction of ciliary axonemes by techniques of membrane perturbation, CaCI2 influx and
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shear, we hypothesized that alterations elicited by HCHO might involve surface membrane components (Hastie et al., 1990b). In previous studies, the membranes were solubilized during the axoneme isolation procedure and thus could not be examined (Hastie et al., 1986). However, with modification of the isolation buffer and method, the ciliary membranes were retained as a fraction that could be investigated further (Hastie et al., 1990a). Biotin was conjugated covalently to reactive sites on the intact epithelium prior to HCHO exposure to identify surface-accessible components. There are some membrane constituents that are inaccessible or unreactive for attachment of label (Hastie et al., 1990a). However, biotin labeling of membranes in epithelial cells has revealed greater numbers of constituents than metabolic labeling methods (Lisanti et al., 1989). The modified method for extraction of cilia, although retaining ciliary membranes, introduces some additional contaminants including the internal, soluble "matrix" proteins of the cilia (Gibbons, 1965), which are released into the membrane fraction by detergent solubilization of the lipid bilayer and do not associate with reconstituted membranes (Hastie et al., 1990a). Thus biotin labeling was considered necessary to distinguish external components of ciliary membranes. Labeling with biotin did not modify subsequent epithelial response to HCHO as evidenced by similar concentration- and time-dependent inhibition of functionally active, biotinylated rabbit tracheal epithelium compared with tracheal explants that were not biotin labeled. Moreover, equivalent decreased yields of extractable cilia from biotin-labeled and nonlabeled epithelium indicate that the HCHO concentrationdependent alterations in the epithelium were not influenced by the presence of biotin label, nor by the modified procedure for isolation of cilia. The HCHO concentration-dependent decline in extractable cilia predictably resulted in reduced membrane material as well. This was demonstrated by significantly decreased biotin-labeled components identifying surface-accessible material in the detergent-soluble membrane + matrix fraction. The total protein content in the membrane + matrix fractions analyzed by SDS-PAGE remained unchanged. However, the majority of these proteins are not biotinylated, indicating that they were probably not surface-accessible components. In addition, many of these same proteins remain in the supernate upon sedimentation of reconstituted membrane vesicles (Hastie et al., 1990a), which is further evidence that these proteins were not membrane constituents. As stated earlier, nonlabeled proteins in the membrane + matrix fraction may consist of the soluble, internal matrix proteins of cilia. Although decreased numbers of extracted cilia would predictably yield less of these matrix proteins as well, decreased permeability of the ciliary membrane would ensure,
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conversely, that more of these proteins remained within the cilia instead of being lost during the cilia isolation procedure. The biotinylated components in the axoneme fractions, by virtue of possessing this covalently linked label, are accessible to the external environment. The majority of these is distinctly different from biotinylated components in the membrane + matrix fractions and resists detergent solubilization, indicating a close association with the internal ciliary axoneme (Hastie et al., 1990a). The total amount of biotin-labeled material in the axoneme fraction remained nearly the same with increasing HCHO concentration. Thus, considered as a proportion of the total protein in the axoneme fraction, biotinylated material increased significantly with increasing HCHO concentration. This trend was supported by significant, concentration-dependent increases in specific biotin-labeled proteins in the axoneme fraction, although other biotinlabeled proteins in this fraction decreased. Thus not all surfaceaccessible components were affected by HCHO in the same way or to the same extent. Casanova-Schmitz et al. (1983) observed decreased extractability of DNA from protein in rat nasal mucosa with increasing HCHO exposure. The increased resistance to normal extraction procedures for cilia from the epithelium and for certain surface-accessible components from association with the axoneme after HCHO exposure may be related phenomena. The pattern of protein migration did not noticeably change with increased HCHO exposure, indicating that molecular weights of the components remained the same. Although the increase of biotinlabeled components in the axoneme fraction might possibly derive from cross-linking of smaller biotin-labeled components, it is more likely that these would appear as new bands, instead of migrating to precisely the same positions as other components already present. Thus covalent linkages between proteins apparently were not formed by HCHO exposure. However, internal methylene bridge formation between amino groups within a protein molecule rather than crosslinking between adjacent molecules (French and Edsall, 1945) may make the epithelial surface components less susceptible to detergent solubilization. In conclusion, we have demonstrated (1) reduced extraction of surface accessible membrane components; (2) increased retention of internal soluble proteins within the cilia, subsequently released into the membrane + matrix fraction; and (3) increased retention of surface accessible components with internal axonemes. Together, these points confirm that components accessible at the epithelial surface were altered to varying degrees by exposure to HCHO, possibly through internal molecular stabilization. Such alterations, if not reversed, may result in other secondary responses leading to loss of cilia and to cell injury and death, which have been observed at higher HCHO concentrations
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or longer exposure periods (Bhalla et al., 1991; Monteiro-Riviere and Popp, 1986; Monticello et al., 1989; Zwart et al., 1988).
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REFERENCES Alexandersson, R., and Hedenstierna, G. 1989. Pulmonary function in wood workers exposed to formaldehyde: A prospective study. Arch. Environ. Health 44:5-11. Bhalla, D. K., Mahavni, V., Nguyen, T., and McClure, T. 1991. Effects of acute exposure to formaldehyde on surface morphology of nasal epithelia in rats. J. Toxicol. Environ. Health 33:171-188. Bradford, M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254. Carson, S., Goldhamer, R., and Carpenter, R. 1966. Mucus transport in the respiratory tract. Am. Rev. Respir. Dis. 93:86-92. Casanova-Schmitz, M., and Heck, H. D'A. 1983. Effects of formaldehyde exposure on the extractability of DNA from proteins in the rat nasal mucosa. Toxicol. Appl. Pharmacol. 70:121-132. Cralley, L. V. 1942. The effect of irritant gases upon the rate of ciliary activity. J. Ind. Hyg. Toxicol. 24:193-198. Dalhamn, T. 1956. Mucous flow and ciliary activity in the trachea of healthy rats and rats exposed to respiratory irritant gases (SO2, N 3 H, HCHO). Acta Physiol. Scand. 36(Suppl. 123):1-161. Eller, P. M., ed. 1984. NIOSH Manual of Analytical Methods, 3d ed., pp. 3500-1-3500-4. Washington, D.C.: Government Printing Office. Publication number 84-100. Falk, H. L., Kotin, P., and Rowlette, W. 1963. The response of mucus-secreting epithelium and mucus to irritants. Ann. N.Y. Acad. Sci. 106:583-608. Fiske, C. H., and Subbarrow, Y. 1925. The colorimetric determination of phosphorus, f. Biol. Chem. 66:375-400. French, D., and Edsall, J. T. 1945. The reactions of formaldehyde with amino acids and proteins. Adv. Protein Chem. 2:277-335. Gibbons, I. R. 1965. Chemical dissection of cilia. Arch. Biol. (Liege) 76:317-352. Hastie, A. T., Dicker, D. T., Hingley, S. I , Kueppers, F., Higgins, M. L., and Weinbaum, G. 1986. Isolation of cilia from porcine tracheal epithelium and extraction of dynein arms. Cell Motil. Cytoskel. 6:25-34. Hastie, A. T., Loegering, D. A., Gleich, G. J., and Kueppers, F. 1987. The effect of purified human eosinophil major basic protein on mammalian ciliary activity. Am. Rev. Respir. Dis. 135:848853. Hastie, A. T., Krantz, M. J., and Colizzo, F. P. 1990a. Identification of surface components of mammalian respiratory tract cilia. Cell Motil. Cytoskel. 17:317-328. Hastie, A. T., Patrick, H., and Fish, J. E. 1990b. Inhibition and recovery of mammalian respiratory ciliary function after formaldehyde exposure. Toxicol. Appl. Pharmacol. 102:282-291. Kensler, C. J., and Battista, S. P. 1966. Chemical and physical factors affecting mammalian ciliary activity. Am. Rev. Respir. Dis. 93:93-102. Krzyanowski, M., Quackenboss, J. J., and Lebowitz, M. D. 1990. Chronic respiratory effect of indoor formaldehyde exposure. Environ. Res. 52:117-125. Lisanti, M., Le Bivic, A., Sargiacomo, M., and Rodriguez-Boulan, E. 1989. Steady-state distribution and biogenesis of endogenous Madin-Darby canine kidney glycoproteins: Evidence for intracellular sorting and polarized cell surface delivery. J. Cell Biol. 109:2117-2127. Maizel, J. V., Jr. 1971. Polyacrylamide gel electrophoresis of viral proteins. Methods Virol. 5:179-246. Monterio-Riviere, N. A., and Popp, J. A. 1986. Ultrastructural evaluation of acute nasal toxicity in the rat respiratory epithelium in response to formaldehyde gas. Fundam. Appl. Toxicol. 6:251262. Monticello, T. M., Morgan, K. T., Everitt, J. I., and Popp, J. A. 1989. Effects of formaldehyde gas on the respiratory tract of rhesus monkeys. Am. J. Pathol. 134:515-527. Morgan, K. T., Gross, E. A., and Patterson, D. L. 1986a. Distribution, progression, and recovery of
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acute formaldehyde-induced inhibition of nasal mucociliary function in F-344 rats. Toxicol. Appl. Pharmacol. 86:448-456. Morgan, K. T., Patterson, D. L., and Gross, E. A. 1986b. Responses of the nasal mucociliary apparatus of F-344 rats to formaldehyde gas. Toxicol. Appl. Pharmacol. 82:1-13. Zwart, A., Woutersen, R. A., Wilmer, J. W. C. M., Spit, B. J., and Feron, V. J. 1988. Cytotoxic and adaptive effects in rat nasal epithelium after 3 day and 13 week exposure to low concentrations of formaldehyde vapour. Toxicology 51:87-99.
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Received August 10, 1991 Accepted November 15, 1991
