- Email: [email protected]
Action of histamine on eustachian tube function
Action of histamine on eustachian tube function BRIAN W. DOWNS, MD, HENRY F. BUTEHORN III, MD, JIRI PRAZMA, MD, PhD, AUSTIN S. ROSE, MD, JOCELYN C. STAMAT, MD, and HAROLD C. PILLSBURY III, MD, Chapel Hill, North Carolina
INTRODUCTION: The role of allergy in eustachian tube dysfunction is controversial. In this study, allergy was simulated by exposure to histamine, and eustachian tube function testing was performed in an experimental rat model. METHODS: Ventilatory function was assessed by measuring passive opening and closing pressures of the eustachian tube after challenge with either transtympanic or intranasal histamine. The mucociliary clearance time of the tubotympanum was assessed by observing dye transport from the middle ear to the nasopharynx after challenge with either transtympanic histamine or control solution. RESULTS: There was a statistically significant increase in passive opening and closing pressures with transtympanic histamine versus intranasal histamine. In addition, mucociliary clearance times of the tubotympanum after transtympanic histamine showed a statistically significant increase when compared with those after transtympanic control solution. CONCLUSIONS: Transtympanic histamine exposure causes eustachian tube dysfunction in the rat by increasing passive opening and closing pressures of the eustachian tube and impairing mucociliary clearance time. (Otolaryngol Head Neck Surg 2001;124:414-20.)
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he role of allergy in eustachian tube dysfunction (ETD) and otitis media with effusion (OME) is controversial. Before the 1960s, anecdotal evidence pointed to a link between allergy and otitis media with effusion. With advances in immunology and a better understanding of From the University of North Carolina at Chapel Hill. Supported by the Deafness Research Foundation Otologic Research Fellowship. Presented at the Annual Meeting of the American Academy of Otolaryngology–Head and Neck Surgery, New Orleans, LA, September 26-29, 1999. Reprint requests: Jiri Prazma, MD, PhD, Division of Otolaryngology–Head and Neck Surgery, UNC School of Medicine, 610 Burnett-Womack Building, CB#7070, Chapel Hill, NC 275997070; e-mail, [email protected]. Copyright © 2001 by the American Academy of Otolaryngology–Head and Neck Surgery Foundation, Inc. 0194-5998/2001/$35.00 + 0 23/1/113943 doi:10.1067/mhn.2001.113943 414
middle ear pathophysiology, the eustachian tube (ET) has been implicated as a target organ for allergy in OME.1 The role of allergy is not universally accepted, however, as the incidence of allergy in OME has been reported to range from near 0 to greater than 80%.2 Further delineation of the role of allergy in ETD is needed to best identify treatment options for otitis media with effusion. The eustachian tube serves 3 roles: ventilation, clearance, and protection of the middle ear.3 One theory holds that the superior aspect of the eustachian tube assists in middle ear ventilation and the inferior aspect assists in clearance of noxious substances via ciliated cells; together these actions help protect the middle ear.4 The breakdown of this protective function predisposes to otitis media. Moreover, the pediatric eustachian tube is shorter and more horizontal in orientation than in the adult, intrinsically impairing its protective function.3,5 In allergy, mediators of inflammation in the nasopharynx block the eustachian tube by causing edema in the tissues surrounding the ET opening, resulting in impairment of ventilation and mucociliary clearance from the middle ear.1 Histamine, one of the best-known mediators, facilitates edema formation by promoting exudation of fluid and proteins from the mucosal vasculature into the extracellular space during an allergic reaction.6 The purpose of this study was to evaluate the eustachian tube function in a rat model of allergy. Allergy was simulated by instilling histamine into the middle ear or nasopharynx. The eustachian tube ventilatory function was assessed using the forced-response test to measure passive opening and closing pressures as described by Flisberg et al.7 In addition, eustachian tube clearance function was assessed using the mucociliary clearance time of the tubotympanum (MCTT),8 in which the transit time of dye from the middle ear to the eustachian tube orifice in the nasopharynx was measured.9 Our hypothesis was that histamine causes acute eustachian tube dysfunction by increasing passive pressures and prolonging MCTTs. MATERIALS AND METHODS Animals This study complies with “The Principles of Laboratory Animal Care” formulated by the National Society for Medical Research and the “Guide for the Care and Use of Laboratory Animals” prepared by the National Academy of Sciences and
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published by the National Institute of Health. In addition, all animals were handled according to Institutional Review Boardapproved protocols. Nonallergic Sprague-Dawley rats, from 250-470 grams, were randomly assigned to receive either transtympanic or intranasal histamine (H-7250; Sigma Chemical Co., St. Louis, MO). Rats undergoing pressure measurements comprised a separate group from those undergoing MCTT measurements. In the pressure group (n = 13), the experimental ear was exposed to 0.01% and 0.1% transtympanic histamine (n = 5), or 1% transtympanic histamine only (n = 5). The remaining 3 rats were exposed to 1% and 10% intranasal histamine administered into the nostril ipsilateral to the ear in which pressures were measured. In the MCTT group (n = 6), the experimental ear was exposed to 1% transtympanic histamine (n = 3) or phosphate-buffered saline (PBS) control solution (n = 3). Surgical Preparation and Perioperative Management
Each rat was anesthetized with 0.1 ml/100 g of a 1:1 mixture of xylazine (20 mg/mL) and ketamine hydrochloride (100 mg/mL). Depth of anesthesia was measured by testing joint pressure receptors. All animals were given 80 µL/min/kg of 300 mOsm Krebs Ringer solution (140 mmol/L Na+, 120 mmol/L Cl-, 5.2 mmol/L K+, 25 mmol/L HCO3-, 1.1 mmol/L Ca2+, 1.2 mmol/L Mg2+, 0.4 mmol/L HPO2-, and 5.6 mmol/L glucose) subcutaneously to maintain blood pressure and renal perfusion during the experiment. A thermistor-controlled heating pad was used to maintain body temperature at 37°C. Measurement of Passive Pressures by the Forced-response Test
Rats were placed in the recumbent position. The tympanic membranes (TMs) were visualized with an operating microscope to rule out preexisting effusion or other pathosis. Two TM incisions were made with a small myringotomy knife, one in the anteroinferior quadrant and one in the posteroinferior quadrant. The posterior myringotomy served as an injection port and the anterior one served as a vent. Baseline passive opening and closing pressures were made via the forced-response test similar to the method first described by Flisberg et al.7 An airtight seal was created in the ear canal with a 6 French latex pediatric foley catheter with a 3 cc balloon (Medline, Mundelein, IL). The balloon was inflated in the external auditory canal to create a closed system. A water manometer was connected to the drainage port of the foley catheter and positive pressure was applied using a rubber sphygmomanometer bulb. Slow uniform pressure was applied to the bulb until the meniscus of the water column changed from convex to concave and the column began descending. This change, or “breaking point,” was considered the passive opening pressure. The
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point at which the water column stopped descending was considered the passive closing pressure. After baseline pressures (time = 0 minutes) were measured, 35 µL of either 0.01% histamine (n = 5), 0.1% histamine (n = 5), or 1% histamine (n = 5) were injected into the middle ear using a 25-gauge needle (the volume of the middle ear space in rats is approximately 35 µL). Pressures were measured at 6, 12, 18, and 24 minutes after injection. Three pressures were taken at each time point to increase accuracy. Raw data were used for statistical analysis, but, for graphic illustration, comparisons were done based on percentage change because each rat had a slightly different pressure at time = 0. The forced-response test was repeated on rats exposed to intranasal histamine. Rats were prepared in a similar fashion, and baseline pressures were obtained. Next, rats were exposed to 12 µL of intranasal histamine via a micropipet in concentrations of 0.01% and 0.1% (n = 1) or 1% and 10% (n = 3). Twelve microliters was the largest intranasal volume that did not cause aspiration and swallowing in rats, thereby compromising ET function tests. Again, 3 pressures each were obtained at 6, 12, 18, and 24 minutes after injection. Data were not statistically analyzed or graphed for the rats receiving the 2 lowest concentrations (0.01% and 0.1%) as no pressure changes were noted. Statistical Analysis
Pressures were analyzed by 2-way repeated-measure analysis of variance with all pairwise multiple comparison procedures (the Student-Newman-Keuls method). Statistical significance was reserved for P values < 0.05. As little effect was seen with the 0.01% concentration, it was considered the control group for statistical purposes. Measurement of Mucociliary Clearance Time (MCTT)
Rats were anesthetized and prepared as described earlier. Deep anesthesia was obtained to prevent swallowing and active opening of the ET. A ventral midline incision was made and a tracheostomy was performed. Using a modified standard nasal speculum as a head-holder device, rats were placed in the supine position to visualize the soft palate. A small midline incision was made in the soft palate just posterior to the posterior border of the hard palate. The eustachian tube orifice was visualized with a miniature hand mirror under the operating microscope. Rats were placed in the right lateral recumbent position and 2 incisions, 1 posteroinferior for injection and 1 anteroinferior for a vent, were made in the left TM with a small myringotomy knife. Next, 35 microliters of either 1% histamine (n = 3) or control PBS solution (n = 3) were injected into the middle ear with a 25-gauge nee-
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Fig 1. Graph shows percent change in opening pressures versus time for all 3 concentrations of transtympanic histamine. Vertical lines represent standard error of the mean. There was a statistically significant increase with 1% histamine at the 18-minute timepoint over all other timepoints with both transtympanic and intranasal histamine.
dle. Approximately 18 minutes later the residual fluid was suctioned from the middle ear and 4 µL of blue dye (Coumassie Brilliant Blue,9 B-0149; Sigma Chemical Co, St Louis, MO) were deposited into the middle ear using a micropipet. A stopwatch was started and rats were returned to the supine position. The nasopharyngeal orifice of the ET was again visualized and the time to appearance of the dye at the orifice was measured. This time was denoted as the mucociliary clearance time.9 Statistical analysis. MCTTs were analyzed using the unpaired t test and statistical significance was reserved for P < 0.05. RESULTS Transtympanic Histamine Passive pressures.In rats exposed to 0.01% histamine, the mean ± standard error of the mean (SEM) passive opening pressures were 58.5 ± 2.5 at baseline, 58.1 ± 2.4 at 6 minutes, 57.1 ± 1.4 at 12 minutes, 53.7 ± 1.8 at 18 minutes, and 55.2 ± 2.1 at 24 minutes. In rats exposed to 0.1% histamine, the mean ± SEM passive opening pressures were 51.3 ± 3.0 at baseline, 62.7 ± 2.7 at 6 minutes, 58.8 ± 2.9 at 12 minutes, 58.2 ± 1.8 at 18 minutes, and 57.0 ± 3.5 at 24 minutes. In rats exposed to 1% histamine, the mean ± SEM passive opening pressures were 49.3 ± 2.5 at baseline, 61.5 ± 6.3 at 6 minutes, 53.1 ± 3.6
at 12 minutes, 71.1 ± 2.8 at 18 minutes, and 53.4 ± 3.3 at 24 minutes. There was a statistically significant increase in pressure with 1% concentration at 18 minutes over 1% at time = 0. In addition, there was a statistically significant increase with 1% solution at 18 minutes when compared with 0.01% and 0.1% at 6, 12, and 24 minutes. The increase in opening pressure at 6 minutes with 0.1% histamine was not statistically significant. In rats exposed to 0.01% histamine, the mean ± SEM passive closing pressures were 20.6 ± 1.3 at baseline, 23.6 ± 3.1 at 6 minutes, 23.4 ± 3.3 at 12 minutes, 19.5 ± 2.2 at 18 minutes, and 17.7 ± 2.5 at 24 minutes. In rats exposed to 0.1% histamine, the mean ± SEM passive closing pressures were 19.0 ± 2.9 at baseline, 26.5 ± 4.0 at 6 minutes, 24.4 ± 3.6 at 12 minutes, 19.9 ± 2.7 at 18 minutes, and 19.0 ± 2.9 at 24 minutes. In rats exposed to 1% histamine, the mean ± SEM passive closing pressures were 13.2 ± 2.0 at baseline, 26.9 ± 3.9 at 6 minutes, 27 ± 2.1 at 12 minutes, 40.3 ± 3.1 at 18 minutes, and 22.1 ± 2.3 at 24 minutes. There was a statistically significant increase in pressure at 1% concentration and 18 minutes over 1% at time = 0 and over the 6, 12, 18, and 24 minute measurements at both 0.01% and 0.1%. In addition, the 1% solution at 18 minutes was statistically significant from the 1% solution at 6, 12, and 24 minutes. Finally the 1% solution showed a statistically
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Fig 2. Graph shows percent change in closing pressures versus time for all 3 concentrations of transtympanic histamine. Vertical lines represent standard error of the mean. There was a statistically significant increase with 1% histamine at the 18-minute timepoint over all other timepoints with both transtympanic and intranasal histamine.
significant increase in pressure at 6 and 12 minutes over the 1% baseline. The increases seen with 0.1% were not statistically significant. The percent change in opening and closing pressures after transtympanic histamine at the 3 concentrations is shown in Figs 1 and 2, respectively. Again, because of slight variations in pressure at time = 0 between rats, results were illustrated on a percentage of change graph. Mucociliary clearance time (MCTT). MCTTs are shown in Fig 3. In rats exposed to transtympanic PBS (control), MCTTs were 55 seconds, 1 minute 16 seconds, and 1 minute 23 seconds. In rats exposed to 1% transtympanic histamine, they were 4 minutes 20 seconds, 15 minutes, and 15 minutes (dye was never visualized at the 15-minute time points but the ET was considered obstructed if no dye appeared after this time).10 MCTTs in rats receiving 1% histamine showed a statistically significant increase over MCTTs in the control group (P = 0.045).
24 minutes. In rats exposed to 10% histamine, the mean ± SEM passive opening pressures were 56.0 ± 3.2 at baseline, 52.7 ± 2.6 at 6 minutes, 53.3 ± 2.9 at 12 minutes, 51.2 ± 0.9 at 18 minutes, and 49.0 ± 2.1 at 24 minutes. None of the opening pressures at either concentration of intranasal histamine showed statistically significant changes, although slight decreases in pressure were noted. In rats exposed to 1% histamine, the mean ± SEM passive closing pressures were 23.1 ± 4.3 at baseline, 14.8 ± 2.3 at 6 minutes, 14.6 ± 3.5 at 12 minutes, 13.0 ± 3.5 at 18 minutes, and 15.1 ± 4.8 at 24 minutes. In rats exposed to 10% histamine, the mean ± SEM passive closing pressures were 21.8 ± 4.9 at baseline, 19.4 ± 2.3 at 6 minutes, 18.3 ± 2.0 at 12 minutes, 16.0 ± 2.5 at 18 minutes, and 15.1 ± 3.4 minutes. None of the closing pressures at either concentration of intranasal histamine showed statistically significant changes, although slight decreases in pressure were again observed.
Intranasal histamine.
Passive opening pressures after 1% transtympanic histamine at 18 minutes showed a statistically significant increase over 1% and 10% intranasal histamine at all time points. In addition, 0.1% transtympanic histamine at 6 minutes showed a statistically significant increase over 1% intranasal histamine at all time points. Similarly, passive closing pressures after 1% transtympanic histamine at 18 minutes showed a statistically sig-
Transtympanic versus intranasal histamine. Passive pressures. Passive opening and closing pressures after the administration of 1% and 10% intranasal histamine are shown in Figs 4 and 5, respectively. In rats exposed to 1% intranasal histamine, the mean ± SEM passive opening pressures were 47.3 ± 5.9 at baseline, 40.2 ± 4.2 at 6 minutes, 36.8 ± 4.6 at 12 minutes, 39.7 ± 3.4 at 18 minutes, and 40.3 ± 3.8 at
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Fig 3. Graph shows MCCT for 1% transtympanic histamine and PBS control solution.
Fig 4. Graph shows opening and closing pressures versus time for 1% histamine administered intranasally. Vertical lines represent standard error of the mean. Changes observed at this concentration were not statistically significant.
nificant increase over 1% and 10% intranasal histamine at all time points. In addition, 0.1% transtympanic histamine at 6 minutes showed a statistically significant increase over 1% intranasal histamine at all time points and 10% intranasal histamine at 18 and 24 minutes.
DISCUSSION
The results of this study indicate that histamine impairs the ventilatory and clearance functions of the rat ET by increasing passive pressures and prolonging MCCTs. After 0.01% histamine instilled transtympanically, no
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Fig 5. Graph shows opening and closing pressures versus time for 10% histamine administered intranasally. Vertical lines represent standard error of the mean. Changes observed at this concentration were not statistically significant.
change was seen in ET opening or closing pressures. After 0.1% transtympanic histamine, slight increases in opening and closing pressures were repeatedly observed, though they were not statistically significant. Finally, 1% transtympanic histamine produced statistically significant increases in both passive opening and closing pressures. In addition, rats receiving 1% histamine transtympanically showed a statistically significant increase in mucociliary clearance time when compared with control rats receiving PBS. In the rats receiving intranasal histamine, however, no increases in passive opening or closing pressures were seen with either 1% or 10% concentration. Our study builds on past work addressing histamine and ET dysfunction.8,11,12 Moreover, recent work in our laboratory represented the first conclusive evidence linking allergen presentation to the middle ear with persistence of effusion in an animal model.13 Past reports have implicated allergy as a cause of ETD after intranasal challenge with seasonal14 and perennial15 allergens, as well as histamine itself.11 However, no previous study has simultaneously compared transtympanic challenge with intranasal challenge, and no previous study has looked at the immediate effects of histamine on mucociliary clearance times. Our goal was to challenge the eustachian tube from the middle ear and from the nasopharynx. In addition, we desired to expand our rat model of otitis media with effusion to further study of eustachian tube function. Several models of intranasal challenge have been published. An atomizer has been used in monkeys to
aerosolize challenge solution,11 and an Eppendorf pipet has been used to administer intranasal challenge solutions to monkeys.12 In addition, nebulization has been used as a mode of intranasal challenge to provoke ETD.16 Hardy et al17 compared intranasal drug delivery systems by focusing on the deposition pattern of spray versus drops. They found that drops of radiolabeled albumin solutions administered to human volunteers spread throughout the nasal cavity and into the pharynx more extensively than spray.17 This method proved successful in our lab when tested using a set volume of blue dye deposited intranasally in the rat. With direct visualization, dye was observed coating the entire nasopharynx, including the nasopharyngeal orifice of the eustachian tube. Doyle et al11 observed increases in passive opening and closing pressures, primarily in juvenile monkeys, with aerosolized intranasal histamine at concentrations as low as 0.5%. Rats in our study that received 1% and 10% intranasal histamine were all juvenile rats, between 260 and 305 g, but no increase in passive opening or closing pressure was seen. This could represent an undefined, species-specific difference in anatomy or function of the eustachian tube. In a separate study, transient positive pressure in the middle ear has been reported within the first 20 minutes after intranasal histamine in monkeys.12 Perhaps our slight decrease in passive opening and closing pressures after intranasal histamine represents this same phenomenon in rats. It is also possible that our testing of passive pressures was
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not sensitive enough to detect ETD with intranasal challenge. Even so, we feel that this is a useful test for characterizing response in an allergic model. Finally, our experiments were done in an acute animal model and do not reflect repeated challenges, which may occur over a prolonged period of time in an allergic child.1 Despite deep anesthesia, the 1% transtympanic histamine solution caused 4 of the 5 rats to swallow repeatedly when taking measurements at 6 and 12 minutes. This was not observed at the lower concentrations, 0.01% and 0.1%. Even with swallowing, we observed a statistically significant difference in passive pressures at 1% histamine administered transtympanically. This swallowing was perhaps the reason for the “bimodal peak” in the 1% transtympanic curve, in contrast to a smoother, more rounded curve which was expected. The swallowing could have been caused by the combination of 1% histamine and the pressure measurement itself, stimulating the tympanic nerve of Jacobson (branch of the glossopharyngeal) and the afferent limb of the gag reflex. Documented neural connections between the middle ear, brain, and eustachian tube could also participate in a reflex-coordinated opening and closing of the eustachian tube18 mediated by histamine. A concentration dependence would explain the presence of this phenomenon at 1% and not at 0.01% or 0.1%. Investigation of this and other neurogenic pathways is currently underway in our laboratory. Impaired mucociliary clearance results in retention of fluid in the middle ear and difficulty clearing effusions, contributing to the chronic nature of OME.9,19 Ciliated cells in the ET beat toward the nasopharynx, with the most active cells on the middle ear side of the tube.20 Dye transport in the ET has been shown to be impaired after exposure to high-dose bacterial endotoxin,9 formalin-fixed Hemophilus influenzae,21 and influenza A virus.10 Esaki et al8 found prolonged MCTTs at 1, 3, and 8 days after injection with histamine and impairment of the in vitro ciliary beat frequency through day 8 as well. Our study represents the first measurement of MCTT within minutes after exposure to histamine, illustrating the effects of immediate hypersensitivity on the eustachian tube. This study describes the different responses of the ET when histamine is administered transtympanically versus intranasally. Nonresponsiveness of the ET to intranasal histamine suggests that, under normal physiologic conditions, histamine cannot gain access to the middle ear against the direction of ciliary beating. Although the role of allergy is not universally accepted, studies continue to show improvement in patients after avoidance of putative allergens.22 Future research should focus on mucociliary clearance and ET tests dur-
ing swallowing, both with positive and negative pressure, to further characterize allergic eustachian tube dysfunction. Additional investigation involving antiallergy therapy is warranted. REFERENCES 1. Bernstein JM. Role of allergy in eustachian tube blockage and otitis media with effusion: a review. Otolaryngol Head Neck Surg 1996;114:562-8. 2. Bernstein JM. The role of IgE-mediated hypersensitivity in the development of otitis media with effusion. Otolaryngol Clin North Am 1992;25:197-211. 3. Bluestone CD. Pathogenesis of otitis media: role of eustachian tube. Pediatr Infect Dis J 1996;15:281-91. 4. Sando I, Takahashi H, Matsune S, et al. Localization of function in the eustachian tube: a hypothesis. Ann Otol Rhinol Laryngol 1994;103:311-4. 5. Procter B. Embryology and anatomy of the eustachian tube. Arch Otolaryngol 1967;86:503-14. 6. Metcalfe DD, Baram D, Mekori YA. Mast cells. Physiol Rev 1997;77:1033-79. 7. Flisberg K, Ingelstedt S, Örtegren U. Controlled “ear aspiration” of air: a “physiological” test of the tubal function. Acta Otolaryngol (Stockh) 1963;S182:35-8. 8. Esaki Y, Ohashi Y, Furuya H, et al. Histamine-induced mucociliary dysfunction and otitis media with effusion. Acta Otolaryngol (Stockh) 1991;S486:116-34. 9. Bakaletz LO, Griffith SR, Lim DJ. Effect of prostaglandin E2 and bacterial endotoxin on the rat of dye transport in the eustachian tube of the chinchilla. Ann Otol Rhinol Laryngol 1989;98:278-82. 10. Park K, Bakaletz LO, Coticchia JM, et al. Effect of influenza A virus on ciliary activity and dye transport function in the chinchilla eustachian tube. Ann Otol Rhinol Laryngol 1993;102:551-8. 11. Doyle WJ, Ingraham AS, Fireman P. The effects of intranasal histamine challenge on eustachian tube function. J Allergy Clin Immunol 1985;76:551-6. 12. Seroky JT, Alper CM, Tabari R, et al. Effects of intranasal challenge with histamine, bradykinin and prostaglandin on middle ear pressure and blood flow in cynomolgus monkeys. Acta Otolaryngol (Stockh) 1995;115:83-7. 13. Labadie RF, Jewett BS, Hart CF, et al. Allergy increases susceptibility to otitis media with effusion in a rat model. Otolaryngol Head Neck Surg 1999;121:687-92. 14. Friedman RA, Doyle WJ, Casselbrandt ML, et al. Immunologicmediated eustachian tube obstruction: a double-blind crossover study. J Allergy Clin Immunol 1983;71:442-7. 15. Skoner DP, Doyle WJ, Chamovitz AH, et al. Eustachian tube obstruction after intranasal challenge with house dust mite. Arch Otolaryngol Head Neck Surg 1986;112:840-2. 16. Skoner DP, Doyle WJ, Fireman P. Eustachian tube obstruction (ETO) after histamine nasal provocation: a doubleblind doseresponse study. J Allergy Clin Immunol 1987;79:27-31. 17. Hardy JG, Lee SW, Wilson G. Intranasal drug delivery by spray and drops. J Pharm Pharmacol 1985;37:294-7. 18. Eden AR. Neural connections between the middle ear, eustachian tube and brain: implications for the reflex control of middle ear aeration. Ann Otol Rhinol Laryngol 1981;90:566-9. 19. Sade J, Eliezer N. Secretory otitis media and the nature of the mucociliary system. Acta Otolaryngol (Stockh) 1970;70:351-7. 20. Ohashi Y, Nakai Y, Koshimo H, et al. Ciliary activity in the in vitro tubotympanum. Arch Otorhinolaryngol 1986;243:317-9. 21. Bakaletz LO, Ohashi Y, Demaria TF, et al. Effect of formalinfixed Hemophilus influenzae and Streptoccus pnemoniae on dye transport by the chinchilla eustachian tube. Acta Otolaryngol (Stockh) 1989;107:235-43. 22. Derebery MJ, Berliner KI. Allergic eustachian tube dysfunction: diagnosis and treatment. Am J Otol 1997;18:160-5.
INTRODUCTION: The role of allergy in eustachian tube dysfunction is controversial. In this study, allergy was simulated by exposure to histamine, and eustachian tube function testing was performed in an experimental rat model. METHODS: Ventilatory function was assessed by measuring passive opening and closing pressures of the eustachian tube after challenge with either transtympanic or intranasal histamine. The mucociliary clearance time of the tubotympanum was assessed by observing dye transport from the middle ear to the nasopharynx after challenge with either transtympanic histamine or control solution. RESULTS: There was a statistically significant increase in passive opening and closing pressures with transtympanic histamine versus intranasal histamine. In addition, mucociliary clearance times of the tubotympanum after transtympanic histamine showed a statistically significant increase when compared with those after transtympanic control solution. CONCLUSIONS: Transtympanic histamine exposure causes eustachian tube dysfunction in the rat by increasing passive opening and closing pressures of the eustachian tube and impairing mucociliary clearance time. (Otolaryngol Head Neck Surg 2001;124:414-20.)
T
he role of allergy in eustachian tube dysfunction (ETD) and otitis media with effusion (OME) is controversial. Before the 1960s, anecdotal evidence pointed to a link between allergy and otitis media with effusion. With advances in immunology and a better understanding of From the University of North Carolina at Chapel Hill. Supported by the Deafness Research Foundation Otologic Research Fellowship. Presented at the Annual Meeting of the American Academy of Otolaryngology–Head and Neck Surgery, New Orleans, LA, September 26-29, 1999. Reprint requests: Jiri Prazma, MD, PhD, Division of Otolaryngology–Head and Neck Surgery, UNC School of Medicine, 610 Burnett-Womack Building, CB#7070, Chapel Hill, NC 275997070; e-mail, [email protected]. Copyright © 2001 by the American Academy of Otolaryngology–Head and Neck Surgery Foundation, Inc. 0194-5998/2001/$35.00 + 0 23/1/113943 doi:10.1067/mhn.2001.113943 414
middle ear pathophysiology, the eustachian tube (ET) has been implicated as a target organ for allergy in OME.1 The role of allergy is not universally accepted, however, as the incidence of allergy in OME has been reported to range from near 0 to greater than 80%.2 Further delineation of the role of allergy in ETD is needed to best identify treatment options for otitis media with effusion. The eustachian tube serves 3 roles: ventilation, clearance, and protection of the middle ear.3 One theory holds that the superior aspect of the eustachian tube assists in middle ear ventilation and the inferior aspect assists in clearance of noxious substances via ciliated cells; together these actions help protect the middle ear.4 The breakdown of this protective function predisposes to otitis media. Moreover, the pediatric eustachian tube is shorter and more horizontal in orientation than in the adult, intrinsically impairing its protective function.3,5 In allergy, mediators of inflammation in the nasopharynx block the eustachian tube by causing edema in the tissues surrounding the ET opening, resulting in impairment of ventilation and mucociliary clearance from the middle ear.1 Histamine, one of the best-known mediators, facilitates edema formation by promoting exudation of fluid and proteins from the mucosal vasculature into the extracellular space during an allergic reaction.6 The purpose of this study was to evaluate the eustachian tube function in a rat model of allergy. Allergy was simulated by instilling histamine into the middle ear or nasopharynx. The eustachian tube ventilatory function was assessed using the forced-response test to measure passive opening and closing pressures as described by Flisberg et al.7 In addition, eustachian tube clearance function was assessed using the mucociliary clearance time of the tubotympanum (MCTT),8 in which the transit time of dye from the middle ear to the eustachian tube orifice in the nasopharynx was measured.9 Our hypothesis was that histamine causes acute eustachian tube dysfunction by increasing passive pressures and prolonging MCTTs. MATERIALS AND METHODS Animals This study complies with “The Principles of Laboratory Animal Care” formulated by the National Society for Medical Research and the “Guide for the Care and Use of Laboratory Animals” prepared by the National Academy of Sciences and
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published by the National Institute of Health. In addition, all animals were handled according to Institutional Review Boardapproved protocols. Nonallergic Sprague-Dawley rats, from 250-470 grams, were randomly assigned to receive either transtympanic or intranasal histamine (H-7250; Sigma Chemical Co., St. Louis, MO). Rats undergoing pressure measurements comprised a separate group from those undergoing MCTT measurements. In the pressure group (n = 13), the experimental ear was exposed to 0.01% and 0.1% transtympanic histamine (n = 5), or 1% transtympanic histamine only (n = 5). The remaining 3 rats were exposed to 1% and 10% intranasal histamine administered into the nostril ipsilateral to the ear in which pressures were measured. In the MCTT group (n = 6), the experimental ear was exposed to 1% transtympanic histamine (n = 3) or phosphate-buffered saline (PBS) control solution (n = 3). Surgical Preparation and Perioperative Management
Each rat was anesthetized with 0.1 ml/100 g of a 1:1 mixture of xylazine (20 mg/mL) and ketamine hydrochloride (100 mg/mL). Depth of anesthesia was measured by testing joint pressure receptors. All animals were given 80 µL/min/kg of 300 mOsm Krebs Ringer solution (140 mmol/L Na+, 120 mmol/L Cl-, 5.2 mmol/L K+, 25 mmol/L HCO3-, 1.1 mmol/L Ca2+, 1.2 mmol/L Mg2+, 0.4 mmol/L HPO2-, and 5.6 mmol/L glucose) subcutaneously to maintain blood pressure and renal perfusion during the experiment. A thermistor-controlled heating pad was used to maintain body temperature at 37°C. Measurement of Passive Pressures by the Forced-response Test
Rats were placed in the recumbent position. The tympanic membranes (TMs) were visualized with an operating microscope to rule out preexisting effusion or other pathosis. Two TM incisions were made with a small myringotomy knife, one in the anteroinferior quadrant and one in the posteroinferior quadrant. The posterior myringotomy served as an injection port and the anterior one served as a vent. Baseline passive opening and closing pressures were made via the forced-response test similar to the method first described by Flisberg et al.7 An airtight seal was created in the ear canal with a 6 French latex pediatric foley catheter with a 3 cc balloon (Medline, Mundelein, IL). The balloon was inflated in the external auditory canal to create a closed system. A water manometer was connected to the drainage port of the foley catheter and positive pressure was applied using a rubber sphygmomanometer bulb. Slow uniform pressure was applied to the bulb until the meniscus of the water column changed from convex to concave and the column began descending. This change, or “breaking point,” was considered the passive opening pressure. The
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point at which the water column stopped descending was considered the passive closing pressure. After baseline pressures (time = 0 minutes) were measured, 35 µL of either 0.01% histamine (n = 5), 0.1% histamine (n = 5), or 1% histamine (n = 5) were injected into the middle ear using a 25-gauge needle (the volume of the middle ear space in rats is approximately 35 µL). Pressures were measured at 6, 12, 18, and 24 minutes after injection. Three pressures were taken at each time point to increase accuracy. Raw data were used for statistical analysis, but, for graphic illustration, comparisons were done based on percentage change because each rat had a slightly different pressure at time = 0. The forced-response test was repeated on rats exposed to intranasal histamine. Rats were prepared in a similar fashion, and baseline pressures were obtained. Next, rats were exposed to 12 µL of intranasal histamine via a micropipet in concentrations of 0.01% and 0.1% (n = 1) or 1% and 10% (n = 3). Twelve microliters was the largest intranasal volume that did not cause aspiration and swallowing in rats, thereby compromising ET function tests. Again, 3 pressures each were obtained at 6, 12, 18, and 24 minutes after injection. Data were not statistically analyzed or graphed for the rats receiving the 2 lowest concentrations (0.01% and 0.1%) as no pressure changes were noted. Statistical Analysis
Pressures were analyzed by 2-way repeated-measure analysis of variance with all pairwise multiple comparison procedures (the Student-Newman-Keuls method). Statistical significance was reserved for P values < 0.05. As little effect was seen with the 0.01% concentration, it was considered the control group for statistical purposes. Measurement of Mucociliary Clearance Time (MCTT)
Rats were anesthetized and prepared as described earlier. Deep anesthesia was obtained to prevent swallowing and active opening of the ET. A ventral midline incision was made and a tracheostomy was performed. Using a modified standard nasal speculum as a head-holder device, rats were placed in the supine position to visualize the soft palate. A small midline incision was made in the soft palate just posterior to the posterior border of the hard palate. The eustachian tube orifice was visualized with a miniature hand mirror under the operating microscope. Rats were placed in the right lateral recumbent position and 2 incisions, 1 posteroinferior for injection and 1 anteroinferior for a vent, were made in the left TM with a small myringotomy knife. Next, 35 microliters of either 1% histamine (n = 3) or control PBS solution (n = 3) were injected into the middle ear with a 25-gauge nee-
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Fig 1. Graph shows percent change in opening pressures versus time for all 3 concentrations of transtympanic histamine. Vertical lines represent standard error of the mean. There was a statistically significant increase with 1% histamine at the 18-minute timepoint over all other timepoints with both transtympanic and intranasal histamine.
dle. Approximately 18 minutes later the residual fluid was suctioned from the middle ear and 4 µL of blue dye (Coumassie Brilliant Blue,9 B-0149; Sigma Chemical Co, St Louis, MO) were deposited into the middle ear using a micropipet. A stopwatch was started and rats were returned to the supine position. The nasopharyngeal orifice of the ET was again visualized and the time to appearance of the dye at the orifice was measured. This time was denoted as the mucociliary clearance time.9 Statistical analysis. MCTTs were analyzed using the unpaired t test and statistical significance was reserved for P < 0.05. RESULTS Transtympanic Histamine Passive pressures.In rats exposed to 0.01% histamine, the mean ± standard error of the mean (SEM) passive opening pressures were 58.5 ± 2.5 at baseline, 58.1 ± 2.4 at 6 minutes, 57.1 ± 1.4 at 12 minutes, 53.7 ± 1.8 at 18 minutes, and 55.2 ± 2.1 at 24 minutes. In rats exposed to 0.1% histamine, the mean ± SEM passive opening pressures were 51.3 ± 3.0 at baseline, 62.7 ± 2.7 at 6 minutes, 58.8 ± 2.9 at 12 minutes, 58.2 ± 1.8 at 18 minutes, and 57.0 ± 3.5 at 24 minutes. In rats exposed to 1% histamine, the mean ± SEM passive opening pressures were 49.3 ± 2.5 at baseline, 61.5 ± 6.3 at 6 minutes, 53.1 ± 3.6
at 12 minutes, 71.1 ± 2.8 at 18 minutes, and 53.4 ± 3.3 at 24 minutes. There was a statistically significant increase in pressure with 1% concentration at 18 minutes over 1% at time = 0. In addition, there was a statistically significant increase with 1% solution at 18 minutes when compared with 0.01% and 0.1% at 6, 12, and 24 minutes. The increase in opening pressure at 6 minutes with 0.1% histamine was not statistically significant. In rats exposed to 0.01% histamine, the mean ± SEM passive closing pressures were 20.6 ± 1.3 at baseline, 23.6 ± 3.1 at 6 minutes, 23.4 ± 3.3 at 12 minutes, 19.5 ± 2.2 at 18 minutes, and 17.7 ± 2.5 at 24 minutes. In rats exposed to 0.1% histamine, the mean ± SEM passive closing pressures were 19.0 ± 2.9 at baseline, 26.5 ± 4.0 at 6 minutes, 24.4 ± 3.6 at 12 minutes, 19.9 ± 2.7 at 18 minutes, and 19.0 ± 2.9 at 24 minutes. In rats exposed to 1% histamine, the mean ± SEM passive closing pressures were 13.2 ± 2.0 at baseline, 26.9 ± 3.9 at 6 minutes, 27 ± 2.1 at 12 minutes, 40.3 ± 3.1 at 18 minutes, and 22.1 ± 2.3 at 24 minutes. There was a statistically significant increase in pressure at 1% concentration and 18 minutes over 1% at time = 0 and over the 6, 12, 18, and 24 minute measurements at both 0.01% and 0.1%. In addition, the 1% solution at 18 minutes was statistically significant from the 1% solution at 6, 12, and 24 minutes. Finally the 1% solution showed a statistically
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Fig 2. Graph shows percent change in closing pressures versus time for all 3 concentrations of transtympanic histamine. Vertical lines represent standard error of the mean. There was a statistically significant increase with 1% histamine at the 18-minute timepoint over all other timepoints with both transtympanic and intranasal histamine.
significant increase in pressure at 6 and 12 minutes over the 1% baseline. The increases seen with 0.1% were not statistically significant. The percent change in opening and closing pressures after transtympanic histamine at the 3 concentrations is shown in Figs 1 and 2, respectively. Again, because of slight variations in pressure at time = 0 between rats, results were illustrated on a percentage of change graph. Mucociliary clearance time (MCTT). MCTTs are shown in Fig 3. In rats exposed to transtympanic PBS (control), MCTTs were 55 seconds, 1 minute 16 seconds, and 1 minute 23 seconds. In rats exposed to 1% transtympanic histamine, they were 4 minutes 20 seconds, 15 minutes, and 15 minutes (dye was never visualized at the 15-minute time points but the ET was considered obstructed if no dye appeared after this time).10 MCTTs in rats receiving 1% histamine showed a statistically significant increase over MCTTs in the control group (P = 0.045).
24 minutes. In rats exposed to 10% histamine, the mean ± SEM passive opening pressures were 56.0 ± 3.2 at baseline, 52.7 ± 2.6 at 6 minutes, 53.3 ± 2.9 at 12 minutes, 51.2 ± 0.9 at 18 minutes, and 49.0 ± 2.1 at 24 minutes. None of the opening pressures at either concentration of intranasal histamine showed statistically significant changes, although slight decreases in pressure were noted. In rats exposed to 1% histamine, the mean ± SEM passive closing pressures were 23.1 ± 4.3 at baseline, 14.8 ± 2.3 at 6 minutes, 14.6 ± 3.5 at 12 minutes, 13.0 ± 3.5 at 18 minutes, and 15.1 ± 4.8 at 24 minutes. In rats exposed to 10% histamine, the mean ± SEM passive closing pressures were 21.8 ± 4.9 at baseline, 19.4 ± 2.3 at 6 minutes, 18.3 ± 2.0 at 12 minutes, 16.0 ± 2.5 at 18 minutes, and 15.1 ± 3.4 minutes. None of the closing pressures at either concentration of intranasal histamine showed statistically significant changes, although slight decreases in pressure were again observed.
Intranasal histamine.
Passive opening pressures after 1% transtympanic histamine at 18 minutes showed a statistically significant increase over 1% and 10% intranasal histamine at all time points. In addition, 0.1% transtympanic histamine at 6 minutes showed a statistically significant increase over 1% intranasal histamine at all time points. Similarly, passive closing pressures after 1% transtympanic histamine at 18 minutes showed a statistically sig-
Transtympanic versus intranasal histamine. Passive pressures. Passive opening and closing pressures after the administration of 1% and 10% intranasal histamine are shown in Figs 4 and 5, respectively. In rats exposed to 1% intranasal histamine, the mean ± SEM passive opening pressures were 47.3 ± 5.9 at baseline, 40.2 ± 4.2 at 6 minutes, 36.8 ± 4.6 at 12 minutes, 39.7 ± 3.4 at 18 minutes, and 40.3 ± 3.8 at
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Fig 3. Graph shows MCCT for 1% transtympanic histamine and PBS control solution.
Fig 4. Graph shows opening and closing pressures versus time for 1% histamine administered intranasally. Vertical lines represent standard error of the mean. Changes observed at this concentration were not statistically significant.
nificant increase over 1% and 10% intranasal histamine at all time points. In addition, 0.1% transtympanic histamine at 6 minutes showed a statistically significant increase over 1% intranasal histamine at all time points and 10% intranasal histamine at 18 and 24 minutes.
DISCUSSION
The results of this study indicate that histamine impairs the ventilatory and clearance functions of the rat ET by increasing passive pressures and prolonging MCCTs. After 0.01% histamine instilled transtympanically, no
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Fig 5. Graph shows opening and closing pressures versus time for 10% histamine administered intranasally. Vertical lines represent standard error of the mean. Changes observed at this concentration were not statistically significant.
change was seen in ET opening or closing pressures. After 0.1% transtympanic histamine, slight increases in opening and closing pressures were repeatedly observed, though they were not statistically significant. Finally, 1% transtympanic histamine produced statistically significant increases in both passive opening and closing pressures. In addition, rats receiving 1% histamine transtympanically showed a statistically significant increase in mucociliary clearance time when compared with control rats receiving PBS. In the rats receiving intranasal histamine, however, no increases in passive opening or closing pressures were seen with either 1% or 10% concentration. Our study builds on past work addressing histamine and ET dysfunction.8,11,12 Moreover, recent work in our laboratory represented the first conclusive evidence linking allergen presentation to the middle ear with persistence of effusion in an animal model.13 Past reports have implicated allergy as a cause of ETD after intranasal challenge with seasonal14 and perennial15 allergens, as well as histamine itself.11 However, no previous study has simultaneously compared transtympanic challenge with intranasal challenge, and no previous study has looked at the immediate effects of histamine on mucociliary clearance times. Our goal was to challenge the eustachian tube from the middle ear and from the nasopharynx. In addition, we desired to expand our rat model of otitis media with effusion to further study of eustachian tube function. Several models of intranasal challenge have been published. An atomizer has been used in monkeys to
aerosolize challenge solution,11 and an Eppendorf pipet has been used to administer intranasal challenge solutions to monkeys.12 In addition, nebulization has been used as a mode of intranasal challenge to provoke ETD.16 Hardy et al17 compared intranasal drug delivery systems by focusing on the deposition pattern of spray versus drops. They found that drops of radiolabeled albumin solutions administered to human volunteers spread throughout the nasal cavity and into the pharynx more extensively than spray.17 This method proved successful in our lab when tested using a set volume of blue dye deposited intranasally in the rat. With direct visualization, dye was observed coating the entire nasopharynx, including the nasopharyngeal orifice of the eustachian tube. Doyle et al11 observed increases in passive opening and closing pressures, primarily in juvenile monkeys, with aerosolized intranasal histamine at concentrations as low as 0.5%. Rats in our study that received 1% and 10% intranasal histamine were all juvenile rats, between 260 and 305 g, but no increase in passive opening or closing pressure was seen. This could represent an undefined, species-specific difference in anatomy or function of the eustachian tube. In a separate study, transient positive pressure in the middle ear has been reported within the first 20 minutes after intranasal histamine in monkeys.12 Perhaps our slight decrease in passive opening and closing pressures after intranasal histamine represents this same phenomenon in rats. It is also possible that our testing of passive pressures was
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not sensitive enough to detect ETD with intranasal challenge. Even so, we feel that this is a useful test for characterizing response in an allergic model. Finally, our experiments were done in an acute animal model and do not reflect repeated challenges, which may occur over a prolonged period of time in an allergic child.1 Despite deep anesthesia, the 1% transtympanic histamine solution caused 4 of the 5 rats to swallow repeatedly when taking measurements at 6 and 12 minutes. This was not observed at the lower concentrations, 0.01% and 0.1%. Even with swallowing, we observed a statistically significant difference in passive pressures at 1% histamine administered transtympanically. This swallowing was perhaps the reason for the “bimodal peak” in the 1% transtympanic curve, in contrast to a smoother, more rounded curve which was expected. The swallowing could have been caused by the combination of 1% histamine and the pressure measurement itself, stimulating the tympanic nerve of Jacobson (branch of the glossopharyngeal) and the afferent limb of the gag reflex. Documented neural connections between the middle ear, brain, and eustachian tube could also participate in a reflex-coordinated opening and closing of the eustachian tube18 mediated by histamine. A concentration dependence would explain the presence of this phenomenon at 1% and not at 0.01% or 0.1%. Investigation of this and other neurogenic pathways is currently underway in our laboratory. Impaired mucociliary clearance results in retention of fluid in the middle ear and difficulty clearing effusions, contributing to the chronic nature of OME.9,19 Ciliated cells in the ET beat toward the nasopharynx, with the most active cells on the middle ear side of the tube.20 Dye transport in the ET has been shown to be impaired after exposure to high-dose bacterial endotoxin,9 formalin-fixed Hemophilus influenzae,21 and influenza A virus.10 Esaki et al8 found prolonged MCTTs at 1, 3, and 8 days after injection with histamine and impairment of the in vitro ciliary beat frequency through day 8 as well. Our study represents the first measurement of MCTT within minutes after exposure to histamine, illustrating the effects of immediate hypersensitivity on the eustachian tube. This study describes the different responses of the ET when histamine is administered transtympanically versus intranasally. Nonresponsiveness of the ET to intranasal histamine suggests that, under normal physiologic conditions, histamine cannot gain access to the middle ear against the direction of ciliary beating. Although the role of allergy is not universally accepted, studies continue to show improvement in patients after avoidance of putative allergens.22 Future research should focus on mucociliary clearance and ET tests dur-
ing swallowing, both with positive and negative pressure, to further characterize allergic eustachian tube dysfunction. Additional investigation involving antiallergy therapy is warranted. REFERENCES 1. Bernstein JM. Role of allergy in eustachian tube blockage and otitis media with effusion: a review. Otolaryngol Head Neck Surg 1996;114:562-8. 2. Bernstein JM. The role of IgE-mediated hypersensitivity in the development of otitis media with effusion. Otolaryngol Clin North Am 1992;25:197-211. 3. Bluestone CD. Pathogenesis of otitis media: role of eustachian tube. Pediatr Infect Dis J 1996;15:281-91. 4. Sando I, Takahashi H, Matsune S, et al. Localization of function in the eustachian tube: a hypothesis. Ann Otol Rhinol Laryngol 1994;103:311-4. 5. Procter B. Embryology and anatomy of the eustachian tube. Arch Otolaryngol 1967;86:503-14. 6. Metcalfe DD, Baram D, Mekori YA. Mast cells. Physiol Rev 1997;77:1033-79. 7. Flisberg K, Ingelstedt S, Örtegren U. Controlled “ear aspiration” of air: a “physiological” test of the tubal function. Acta Otolaryngol (Stockh) 1963;S182:35-8. 8. Esaki Y, Ohashi Y, Furuya H, et al. Histamine-induced mucociliary dysfunction and otitis media with effusion. Acta Otolaryngol (Stockh) 1991;S486:116-34. 9. Bakaletz LO, Griffith SR, Lim DJ. Effect of prostaglandin E2 and bacterial endotoxin on the rat of dye transport in the eustachian tube of the chinchilla. Ann Otol Rhinol Laryngol 1989;98:278-82. 10. Park K, Bakaletz LO, Coticchia JM, et al. Effect of influenza A virus on ciliary activity and dye transport function in the chinchilla eustachian tube. Ann Otol Rhinol Laryngol 1993;102:551-8. 11. Doyle WJ, Ingraham AS, Fireman P. The effects of intranasal histamine challenge on eustachian tube function. J Allergy Clin Immunol 1985;76:551-6. 12. Seroky JT, Alper CM, Tabari R, et al. Effects of intranasal challenge with histamine, bradykinin and prostaglandin on middle ear pressure and blood flow in cynomolgus monkeys. Acta Otolaryngol (Stockh) 1995;115:83-7. 13. Labadie RF, Jewett BS, Hart CF, et al. Allergy increases susceptibility to otitis media with effusion in a rat model. Otolaryngol Head Neck Surg 1999;121:687-92. 14. Friedman RA, Doyle WJ, Casselbrandt ML, et al. Immunologicmediated eustachian tube obstruction: a double-blind crossover study. J Allergy Clin Immunol 1983;71:442-7. 15. Skoner DP, Doyle WJ, Chamovitz AH, et al. Eustachian tube obstruction after intranasal challenge with house dust mite. Arch Otolaryngol Head Neck Surg 1986;112:840-2. 16. Skoner DP, Doyle WJ, Fireman P. Eustachian tube obstruction (ETO) after histamine nasal provocation: a doubleblind doseresponse study. J Allergy Clin Immunol 1987;79:27-31. 17. Hardy JG, Lee SW, Wilson G. Intranasal drug delivery by spray and drops. J Pharm Pharmacol 1985;37:294-7. 18. Eden AR. Neural connections between the middle ear, eustachian tube and brain: implications for the reflex control of middle ear aeration. Ann Otol Rhinol Laryngol 1981;90:566-9. 19. Sade J, Eliezer N. Secretory otitis media and the nature of the mucociliary system. Acta Otolaryngol (Stockh) 1970;70:351-7. 20. Ohashi Y, Nakai Y, Koshimo H, et al. Ciliary activity in the in vitro tubotympanum. Arch Otorhinolaryngol 1986;243:317-9. 21. Bakaletz LO, Ohashi Y, Demaria TF, et al. Effect of formalinfixed Hemophilus influenzae and Streptoccus pnemoniae on dye transport by the chinchilla eustachian tube. Acta Otolaryngol (Stockh) 1989;107:235-43. 22. Derebery MJ, Berliner KI. Allergic eustachian tube dysfunction: diagnosis and treatment. Am J Otol 1997;18:160-5.