Asthma Research Group and Firestone Institute of Respiratory Health, Department of Medicine, McMaster University and St. Joseph's Hospital, Hamilton, Ontario, Canada L8N 4A6
![]() |
ABSTRACT |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Intranasal instillation techniques are used to deliver various substances to the upper and lower respiratory tract (URT and LRT) in mice. Here, we quantify the relative distribution achieved with intranasal delivery of a nonabsorbable tracer, 99mTc-labeled sulfide-colloid. Relative distribution was determined by killing mice after instillation and quantifying the radioactivity in dissected tissues using gamma scintigraphy. A significant effect of delivery volume on relative distribution was observed when animals were killed 5 min after instillation delivered under gas anesthesia. With a delivery volume of 5 µl, no radiation was detected in the LRT; this increased to a maximum of 55.7 ± 2.5% distribution to the LRT when 50 µl were delivered. The majority of radiation not detected in the LRT was found in the URT. Over the course of the following 1 h, radiation in the LRT remained constant, while that in the URT decreased and appeared in the gastrointestinal tract. Instillation of 25 µl into anesthetized mice resulted in 30.1 ± 6.9% distribution to the LRT, while only 5.3 ± 1.5% (P < 0.05) of the same volume was detected in the LRT of awake mice. Varying the body position of mice did not affect relative distribution. When using intranasal instillation, the relative distribution between the URT and LRT and the gastrointestinal tract is heavily influenced by delivery volume and level of anesthesia.
gamma scintigraphy; topical treatment; upper respiratory tract; lower respiratory tract
![]() |
INTRODUCTION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
THE ADMINISTRATION OF SUBSTANCES to mice by the intranasal route is an effective, noninvasive technique employed for the delivery of allergens (3, 4, 26), drugs or gene therapy (4, 20), immunotherapy (1, 7, 11), and pathogens (12, 16, 18) to the upper and lower respiratory tracts (URT and LRT). In published studies, volumes of substances intranasally instilled into mice range from 5 µl (22) to 100 µl (3), with little justification for the chosen volumes. Intranasal delivery is often carried out after intraperitoneal (9, 19) or inhalation (5, 14, 15) anesthesia but has also been performed with fully awake mice (1, 6, 17). The position of the mouse during intranasal delivery has also varied between studies, with horizontal (13) and head-down supine (12) positions having been used. The specific delivery protocols used in these studies are thought to influence the relative distribution of the delivered substance to the URT, LRT, and gut. However, to our knowledge, very little published information is available describing the distribution of intranasally delivered substances or how the distribution can be influenced by delivery techniques.
Preliminary studies by Tsuyuki et al. (23) have shown that 75% of a 50-µl dose of intranasally administered Evans blue dye is deposited in the airways, with no dye detectable in the esophagus or stomach. Eyles et al. (7) reported that 48% of a 50-µl dose of intranasally instilled 7-µm-diameter 46Sc-labeled styrene-divinyl benzene microspheres was evident in the lungs 15 min after challenge, while Takafuji et al. (21) found that only 19% of a 25-µl dose of 125I-labeled ovalbumin was detected in the lungs at this same time. Similar results have been observed when detection assays were performed 60 min after intranasal administration (8, 21). However, in none of these studies were delivery techniques compared. Thus, although these studies support the use of intranasal delivery as a means for depositing substances in the airway, there is a need for determining the influence of factors such as instillation volume, time, body position, and anesthesia on this distribution. This knowledge would allow investigators to adopt the technique that results in the optimal delivery to the organ of interest while keeping the amount of agents in other tissues to a minimum. Ideally, this information would also allow some degree of quantification of the delivered substances.
By delivering a radioactive tracer to the mouse, we are able to monitor where intranasally instilled substances are distributed, allowing for a better understanding of which parts of the mouse are being exposed to similarly administered allergens, drugs, gene therapies, immunotherapies, and pathogens. In these experiments, a nonabsorbable tracer, 99mTc-labeled sulfide-colloid (99mTc-SC) was used, inasmuch as it has been shown to be a successful tracer for determining the absolute dose of radioactivity deposited in the lungs of small animals (10).
The aims of the present study are to characterize the relative distribution after intranasal instillation and to examine the effects of instillation volume, time, body position, and anesthesia on the relative distribution of substances in mice.
![]() |
METHODS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Animals
Female BALB/c mice were purchased (Harlan Sprague Dawley, Indianapolis, IN) at 8-10 wk of age and housed under specific-pathogen-free conditions. Mice were studied in groups of six. For each group, a single volume of 99mTc-SC was delivered intranasally dropwise to the nares, and the distribution of the substance was assessed via autoradiography of dissected tissue. Details of the 99mTc-SC delivery were altered between groups so that the effects of delivered volume, time, body position, and anesthesia could be assessed. All procedures were reviewed and approved by the Animal Research Ethics Board at McMaster University.99mTc-SC Preparation
99mTc-SC (148 MBq/ml; >75% between 0.2 and 0.8 µm and <5% >0.8 µm; >95% radiopurity) was prepared by the Nuclear Pharmacy at McMaster University Medical Centre according to sulfide colloid kit protocols. Briefly, sodium thiosulfate (0.8 mg/ml), potassium perrhenate (0.13 mg/ml), gelatin USP (1.06 mg/ml), and 1 N HCl (0.07 ml) were combined with sodium pertechnetate (740 MBq/ml) and boiled in sterile water. A phosphate buffer consisting of sodium phosphate dibasic (24.8 mg/ml) and sodium phosphate monobasic (2.13 mg/ml) was added to the mixture, which was cooled to room temperature before a further dilution of 99mTc-SC was made in a pipette tube with a second addition of phosphate buffer solution.99mTc-SC Delivery
Volume study. Animals were lightly anesthetized by inhaled isoflurane (Abbott Laboratories, Montreal, PQ, Canada), and a single volume (5, 10, 15, 20, 25, 50, or 75 µl) of 99mTc-SC was intranasally delivered dropwise to the nares using a pipetman (model P20 or P200, Gilson) while the mouse was in a supine position. At the time of dissection, mice were anesthetized by intraperitoneal injection of 250 µl of ketamine (0.44 mg/ml; Bimeda-MTC, Cambridge, ON, Canada)-xylazine (6.3 mg/ml; Bayer, Etobicoke, ON, Canada) in normal saline. The time between 99mTc-SC administration and tissue dissection was 5 ± 2 min.
Time course study. Animals were lightly anesthetized using inhaled isoflurane, and a single dose of 10, 25, or 75 µl of 99mTc-SC was intranasally delivered. Mice in the 5-min group were then anesthetized by intraperitoneal injection of 250 µl of ketamine-xylazine. The 15-min group was allowed to recover after intranasal administration of 10, 25, or 75 µl of 99mTc-SC and anesthetized by intraperitoneal injection of 250 µl of ketamine-xylazine at 10 min after intranasal delivery. Likewise, the 60-min group was permitted to recover before intraperitoneal injection of 250 µl of ketamine-xylazine at 55 min after intranasal delivery.
Body position study. Animals were lightly anesthetized with isoflurane and positioned in an ~60° inclined or 60° declined supine position by securing the fore- and hindpaws with Velcro to a small board oriented at the desired angle. Mice were briefly exposed to isoflurane to maintain a level of light anesthesia before an intranasal challenge with a single dose of 25 µl of 99mTc-SC was delivered slowly into the nares. Before dissection, mice were again briefly exposed to the inhalation anesthesia and then further anesthetized by intraperitoneal injection of 250 µl of ketamine-xylazine in normal saline. Animals were maintained in the desired position for ~3 min after intranasal delivery, and tissue was dissected 5 ± 2 min after intranasal delivery.
Anesthesia study. Animals were lightly anesthetized with isoflurane, inhaled halothane (MTC-Pharmaceuticals, Cambridge, ON, Canada), or intraperitoneal 2,2,2-tribromoethanol (Avertin, Sigma-Aldrich, Oakville, ON, Canada; 240 mg/ml) or left unanesthetized before intranasal administration of 25 µl of 99mTc-SC. Mice that were anesthetized with the inhalation anesthetics for 99mTc-SC delivery were briefly re-exposed to achieve a level of light anesthesia and then anesthetized by intraperitoneal injection with 250 µl of ketamine-xylazine before tissue dissection. The group anesthetized with 2,2,2-tribromoethanol received no further treatment, while the unanesthetized group was also anesthetized with ketamine-xylazine before tissue dissection. The time between 99mTc-SC administration and tissue dissection was 5 ± 2 min.
Tissue Separation
Tissues were dissected within 2 min of the desired time (5, 15, or 60 min). The abdomen was first dissected for isolation and removal of the stomach and intestine. The trachea was then exposed, and the rib cage was separated along the midline of the sternum. The heart was isolated and discarded. The trachea-esophagus combination was cut at the level of the epiglottis and removed along with the intact lungs. The esophagus was separated from the trachea and lungs, and the lungs were then dissected from the trachea. Finally, the head of the mouse, reflecting the URT, was removed at the base of the skull and top of the cervical spine. All dissected tissue samples, once dissected, were immediately placed in individual polypropylene test tubes for auto-gamma counting.Autoradiography
The proportion of radioactivity in each tissue sample was quantified using a Cobra II auto-gamma counter (Canberra-Packard Canada, Mississauga, ON, Canada). Counts were determined using an energy window of 140 keV ± 15% for the 99mTc isotope (25). Counts for each sample were expressed in counts per minute (cpm) and adjusted by subtracting radioactive counts for the mean of three blank tubes. Final counts for all tissues were corrected to a common time using a 6.02-h half-life for the 99mTc isotope. The radioactivity remaining in the mouse carcasses after dissection of all relevant tissues was quantified using a dose calibrator (model CRC-12, Capintec, Pittsburgh, PA) and expressed in units of MBq. This was necessary, inasmuch as the carcass could not fit into the polypropylene test tubes used for the Cobra II auto-gamma counter. A linear equation was determined by performing a linear regression between the quantity of radioactivity recovered in the head tissue samples as measured using the Capintec radioisotope calibrator (MBq) and the amount of radioactivity in the head tissue sample as calculated by the Cobra II auto-gamma counter (cpm), corrected to the same zero time. This equation was then used to convert the counts in the mouse carcass to counts corresponding to those from the auto-gamma counter (cpm). In preliminary studies, we were able to account for 85-100% of the delivered radiation by comparing the tissue samples with standards containing the same quantities of radioactivity that had been administered to the mice.Statistical and Data Analysis
All values of radioactivity are expressed as a percentage of the total recovered 99mTc-SC detected from each mouse. Values are arithmetic means ± SE. Statistical comparison between groups was made using a mixed-model analysis of variance, with tissue type as a repeated factor and 99mTc-SC delivery technique as a nonrepeated factor. Separate analyses were performed for the volume, time course, anesthesia, and body position experiments. Specific means were compared using Student-Newman-Keuls post hoc analysis when indicated by a significant result from the overall analysis of variance.The volume-dose relationship for each tissue type was established by
fitting the data to the following relationship
![]() |
![]() |
RESULTS |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Volume Study
Altering the volume of 99mTc-SC instilled into mouse nares in this study affected the distribution of radioactivity in the various tissues (Fig. 1). Statistically significant volume-distribution relationships were observed for the lungs (r2 = 0.93), head (r2 = 0.95), and carcass (r2 = 0.60; Fig. 2).
|
|
No radioactivity was detected in the lungs after instillation of 5 µl into the nares. Radioactivity in the lungs increased to 16.4 ± 1.6% after instillation of 10 µl (P < 0.05). The relative distribution to the lungs continued to increase with increasing instilled volume up to 50 µl. At this volume, 55.7 ± 2.5% of the delivered radioactivity was detected in the lungs. No further increase in the relative distribution to the lungs occurred with the instillation of 75 µl (54.7 ± 1.9%, P > 0.05).
Minimal amounts of radioactivity were detected in the stomach 5 min after instillation with all volumes. After instillation of 5 µl, there was no detectable radiation in the stomach. Although 6.6 ± 3.6% of the total radiation was recovered from the stomach of the 25-µl group, there was no significant difference in the relative amount of recovered radioactivity in the stomach between any of the instilled volumes.
The proportion of radioactivity measured in the head 5 min after intranasal delivery was reciprocal to that observed in the lungs. Instillation of 5 µl resulted in 98.7 ± 0.3% of the radioactivity being detected in the head. This decreased to 75.3 ± 2.3% (P < 0.05) after instillation of 10 µl and continued to decrease as the instilled volume increased. After instillation of 50 µl, 26.0 ± 1.9% of the radioactivity was detected in the head. No further decrease in distribution to the head occurred with the instillation of 75 µl (26.9 ± 0.6%, P > 0.05).
The relative distribution of radioactivity in the carcass exhibited results similar to those in the lungs, with increasing percentages of recovered radioactivity in response to increasing volumes of instilled 99mTc-SC (P < 0.05). The percentages of recovered radioactivity in the carcass ranged from 0.7 ± 0.3% (5 µl) to 13.5 ± 1.1% (75 µl).
Time Course Study
Except in the 10-µl group, the volume of radioactivity tended to persist in the lungs, while that in the head appeared to drain into the stomach and intestine over the 60-min measurement period (Fig. 3). Radioactivity recovered from the lungs of mice instilled with 10 µl varied from 21.9 ± 3.5% (5 min) to 6.9 ± 1.9% (60 min, P < 0.05). Instillation of 25 and 75 µl resulted in a relative distribution of 38.8 ± 2.4 and 56.7 ± 5.2% at 5 min; this did not change appreciably over the next 1 h. However, the percentage of radioactivity recovered in the head decreased during the 1 h after instillation in all volume groups (Fig. 3). The relative distribution decreased from 71.7 ± 3.3% (5 min) to 15.1 ± 0.9% (60 min, P < 0.05) with instillation of 10 µl, from 49.3 ± 2.9% (5 min) to 8.3 ± 0.8% (60 min, P < 0.05) with instillation of 25 µl, and from 22.5 ± 1.5% (5 min) to 5.9 ± 1.7% (60 min, P < 0.05) with instillation of 75 µl into the mouse nares.
|
The decreasing relative distribution of radioactivity in the head with time was associated with increasing amounts distributed to the stomach and intestine. The percentage of radioactivity recovered in the stomach was significantly greater in all groups at 15 min than at 5 min. Minimal quantities of radioactivity were detected at 5 min, while 42.2 ± 5.2% (10 µl), 24.1 ± 1.7% (25 µl,) and 20.9 ± 3.5% (75 µl) were recovered 15 min after instillation. At 60 min, only 10 µl resulted in a significant decrease to 6.9 ± 1.9% of recovered radioactivity in the stomach compared with that recovered at 15 min.
There was no radioactivity in the intestine at 5 min in any volume group. The percentage of radioactivity recovered increased to 21.9 ± 5.3% (10 µl), 15.4 ± 3.8% (25 µl), and 4.9 ± 2.6% (75 µl) at 15 min (P < 0.05), further increasing to 72.0 ± 2.3% (10 µl), 35.2 ± 7.0% (25 µl), and 21.8 ± 7.0% (75 µl) at 60 min (P < 0.05). This increase in the percentage of recovered radioactivity in the intestine corresponded directly with the decrease in radioactivity detected in the head from 5 to 60 min and, subsequently, in the stomach from 15 to 60 min.
Body Position Study
The nose-up and nose-down supine positions did not result in any significant difference in the relative distribution of radioactivity compared with the control (i.e., horizontal supine) position (Fig. 4). Percentages of radioactivity recovered in the lungs of mice in nose-up and nose-down orientations were 37.3 ± 3.7 and 39.0 ± 3.4%, respectively, which were not significantly different from 38.8 ± 2.4% in the control group.
|
Similarly, although there was a difference in the relative distribution of radioactivity in the stomach between the nose-up and nose-down groups (1.0 ± 0.8 and 7.0 ± 2.3%, respectively) with respect to the control group (0.6 ± 0.3%), the differences were not statistically significant.
The relative distribution of radioactivity in the head of the respective groups ranged from 37.2 ± 1.5 to 45.9 ± 1.7%, while the percentage of recovered radioactivity in the carcass ranged from 8.8 ± 2.4 to 10.7 ± 3.0%, but neither tissue showed statistically significant differences between groups.
Anesthesia Study
The relative distribution of radioactivity in the tissues was not statistically different between any of the anesthetized mice. There were significant differences in the percentages of recovered radioactivity from the lungs and the stomach of the unanesthetized mice compared with all anesthetized mice (Fig. 5).
|
The quantity of radioactivity recovered from the lungs of the anesthetized mice was 30.1 ± 6.9% (2,2,2-tribromoethanol) and 38.8 ± 2.4% (isoflurane), while that in the unanesthetized mice was 5.3 ± 1.5% (P < 0.05).
Minimal quantities of radioactivity were detected in the stomach of the anesthetized mice. In mice anesthetized with 2,2,2-tribromoethanol or halothane, 0.4 ± 0.3 or 1.1 ± 0.7% was detected, respectively, while 20.6 ± 3.9% (P < 0.05) was measured in the awake mice. Anesthesia did not significantly affect the relative distribution of radioactivity in any other tissues (Fig. 5).
![]() |
DISCUSSION |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We have shown that instillation volume can influence the relative distribution of substances instilled into nares of mice, confirming and extending observations of Eyles et al. (7). We have further observed that the presence, but not the type, of anesthesia also influences this distribution, while body position at the time of instillation does not appear to have an effect. These findings will be useful in designing experiments to maximally distribute substances to the URT or LRT in mice.
The nasal instillation of increasing volumes of fluid in mice resulted in greater relative dosing to the lungs. Curve-fitting analysis suggests that the maximum distribution to the lungs, ~50-60%, might be achieved with a delivery volume of 35 µl. These findings are supported by those of Eyles et al. (7), who observed greater relative distribution to the lungs with 50- than with 10-µl intranasal instillation.
Our finding, that ~55% of the administered dose was found in the LRT, agrees with that of Eyles et al. (7), who showed that 48% of 50 µl of intranasally instilled 7-µm-diameter 46Sc-labeled styrene-divinyl benzene microspheres was evident in the lungs 15 min after instillation. Tsuyuki et al. (23), using similar delivery techniques, observed that 75% of 50 µl of intranasally instilled Evans blue dye deposited in the airways. Methods used to quantify dye in that study were not given. Our observation that there was no radioactivity in the lungs after the 5-µl instillation is, to our knowledge, novel and important, inasmuch as this volume can be used for delivery to the URT with minimal exposure to the LRT.
Initial interpretation of our findings may be that clearance of 99mTc-SC from the LRT during the 1 h after instillation occurred only with an instilled volume of 10 µl. However, one can calculate the approximate volume of radioactivity cleared from the LRT (e.g., with 10 µl, a change from 21.9 to 5.0% over 60 min represents a clearance of 1.7 µl). The corresponding clearances after instillation of 25 and 75 µl were 0.2 and 2.5 µl, respectively. Thus, although there was variability in these measurements, similar volumes were cleared regardless of instillation volume.
Radioactivity not accounted for in the lungs at 5 min could be found in the head (URT), from which the radioactivity rapidly drained to the gastrointestinal (GI) tract during the remaining 1 h. The rapid initial appearance of radioactivity in the lungs, with no subsequent increase, despite the relatively large quantities of radioactivity remaining in the head, suggests that aspiration, rather than aerosolization, of fluids accounted for the initial distribution to the lungs. The alternative would be that fluid in the URT was nebulized and distributed to the LRT. However, if this mechanism were in place, we would have expected to see a progressive increase in the amount of radioactivity distributed to the LRT. For example, in the 75-µl group, 22.5% of the radiation remained in the URT 5 min after instillation, which could potentially have been nebulized.
Over time, we observed radiation in the GI tract with all intranasal instillation volumes. This does not support delivering small volumes as a means to localize distribution to the respiratory tract, as has been claimed (2). Furthermore, our results are in disagreement with those of Tsuyuki et al. (23), who did not detect any Evans blue dye in the esophagus or stomach after intranasal instillation of 50 µl. Possibly these measurements were made immediately after instillation, when distribution to the GI tract is still relatively small.
Although we have provided a time course of the movement of a nonabsorbable tracer after intranasal instillation, it is important to note that the actual site of absorption will be influenced by the physical properties of the delivered substances as well as the local perfusion (24). For example, although we have observed that 99mTc-SC drained from the URT to the GI tract between 5 and 60 min, uptake of lipophilic substances would be greater across the nasal mucosa during this period than uptake of hydrophilic substances.
The position of mice during and immediately after intranasal instillation did not appear to affect the relative distribution of the tracer. Therefore, adoption of altered orientation to optimize delivery to the GI tract or the URT or LRT does not appear warranted.
Type of anesthesia does not effect the distribution of instilled substances. It was anticipated that deeper anesthesia, (intraperitoneal ketamine-xylazine) may have resulted in greater aspiration and, hence, greater distribution to the LRT; however, this was not the case. We suggest that inhalation anesthesia allows for quicker recovery time and offers obvious advantages for animal care.
We observed that intranasal instillation of substances to awake mice resulted in less distribution to the lungs than in anesthetized mice. These results suggest that lack of anesthesia during intranasal instillation reduces but does not eliminate aspiration.
The distribution of radioactivity in the carcass immediately after administration of 99mTc-SC increases with increasing administered volumes (as it does in the head and lung). This increasing distribution of radioactivity in the carcass is hypothesized to result from seepage from tissues during dissection. We believe that the most likely source of the radioactivity in the carcass was leakage from the URT during the time after dissection of the esophagus before decapitation.
In conclusion, instillation volume, as well as presence of anesthesia,
affects the relative distribution of intranasally instilled substances,
while body position and type of anesthesia do not. For optimal delivery
to the LRT, a volume of 35 µl should be delivered to anesthetized
mice. Relative distribution to the URT is best achieved after
intranasal instillation of 5 µl to anesthetized mice. Sparing of the
LRT may be increased but not eliminated in awake mice.
![]() |
ACKNOWLEDGEMENTS |
---|
We thank F. Rashid (Dept. of Nuclear Medicine, McMaster University) and R. Ellis for suggestions and assistance.
![]() |
FOOTNOTES |
---|
This work was supported by a grant from the Father Sean O'Sullivan Research Centre at St. Joseph's Hospital and the Medical Research Council (Canada). Access to nuclear medicine services was made available through funding from the Canadian Cystic Fibrosis Foundation (SPARX). P. M. O'Byrne is a Medical Research Council (Canada) senior scientist. Radioisotopes were generously provided by the Regional Radiopharmacy, Hamilton Health Science.
Address for reprint requests and other correspondence: M. D. Inman, Asthma Research Group, Firestone Regional Chest & Allergy Unit, Rm. 113, 50 Charlton Ave. E, Hamilton, ON, Canada L8N 4A6 (E-mail: inmanma{at}mcmaster.ca).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
10.1152/ajplung.00173.2001
Received 17 May 2001; accepted in final form 6 November 2001.
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
1.
Barackman, JD,
Ott G,
and
O'Hagan DT.
Intranasal immunization of mice with influenza vaccine in combination with the adjuvant LT-R72 induces potent mucosal and serum immunity which is stronger than that with traditional intramuscular immunization.
Infect Immun
67:
4276-4279,
1999
2.
Campbell, EM,
Charo IE,
Kunkel SL,
Strieter RM,
Boring L,
Gosling J,
and
Lucaks NW.
Monocyte chemoattractant protein-1 mediates cockroach allergen-induced bronchial hyperreactivity in normal but not CCR2 /
mice: the role of the mast cells.
J Immunol
163:
2160-2167,
1999
3.
Charpoval, S,
Nabozny G,
Marietta EV,
Raymond EL,
Krco CJ,
and
Andrivet P.
Short ragweed allergen induces eosinophilic lung disease in HLA-DQ transgenic mice.
J Clin Invest
103:
1707-1717,
1999
4.
Chun, S,
Daheshia M,
Lee S,
Eo SK,
and
Touse BT.
Distribution fate and mechanism of immune modulation following mucosal delivery of plasmid DNA encoding IL-10.
J Immunol
163:
2393-2402,
1999
5.
De Bie, JJ,
Henricks PA,
Cruikshank WW,
Hofman G,
van Nijkamp FP,
and
Oosterhout AJ.
Effect of interleukin-16-blocking peptide on parameters of allergic asthma in a murine model.
Eur J Pharmacol
383:
189-196,
1999[ISI][Medline].
6.
DeHann, L,
Holtrop M,
Verweij WR,
Agsteribbe E,
and
Wilschit J.
Mucosal immunogenicity and adjuvant activity of the recombinant A subunit of the Escherichia coli, heat-liable enterotoxin.
Immunology
97:
706-713,
1999[ISI][Medline].
7.
Eyles, JE,
Williamson ED,
and
Alpar HO.
Immunological responses to nasal delivery of free and encapsulated tetanus toxoid: studies on the effect of vehicle volume.
Int J Pharmaceut
189:
75-79,
2000[ISI].
8.
Fenton, RJ,
Morley PJ,
Owens IJ,
Gower D,
Parry S,
Crossman L,
and
Wong T.
Chemoprophylaxis of influenza A virus infections, with single dose of Zanamivir, demonstrates that Zanamivir is cleared slowly from the respiratory tract.
Antimicrob Agents Chemother
43:
2642-2647,
1999
9.
Flano, E,
Woodland DL,
and
Blackman MA.
Requirements for CD4+ T cells in V beta 4+CD8+ T cell activation associated with latent murine gamma herpes virus infection.
J Immunol
163:
3403-3408,
1999
10.
Fok, TF,
Al-Essa M,
Kirpilani H,
Monkman S,
Brown BW, Jr,
Coates G,
and
Dolovich M.
Estimation of pulmonary deposition of aerosol using gamma scintigraphy.
J Aerosol Med
12:
9-15,
1999[ISI][Medline].
11.
Fukushima, A,
Yoo YC,
Yoshimatsu K,
Matsuzawa K,
Tamura M,
Tono-Oko S,
Taniguchi K,
Urasawa S,
Arikawa J,
and
Azuma I.
Effect of MDP-Lys (L18) as a mucosal immunoadjuvant on protection of mucosal infections by Sendai virus and rotavirus.
Vaccine
14:
485-491,
1996[ISI][Medline].
12.
Hvalbye, BKR,
Aaberge IS,
Lovik M,
and
Haneberg B.
Intranasal immunization with heat-inactivated Streptococcus pneumoniae protects mice against systemic pneumococcal infection.
Infect Immun
67:
4320-4325,
1999
13.
Inman, MD,
Ellis R,
Wattie J,
Denburg JA,
and
O'Byrne PM.
Allergen-induced increases in airway responsiveness, airway eosinophilia and bone marrow eosinophil progenitors in mice.
Am J Respir Cell Mol Biol
21:
473-479,
1999
14.
John-Schmid, B,
Wiedermann U,
Bohle B,
Repa A,
Kraft D,
and
Ebnex C.
Oligodeoxynucleotides containing CpG motifs modulate the allergic Th2 responses of BALB/c mice to Bet V1, the major birch pollen allergen.
J Allergy Clin Immunol
104:
1015-1023,
1999[ISI][Medline].
15.
Kobayashi, H,
Horner AA,
Takabayashi K,
Nguyen MD,
Cinman N,
and
Raz E.
Immunostimulatory DNA pre-priming: a novel approach for prolonged Th 1-biased immunity.
Cell Immunol
198:
69-75,
1999[ISI][Medline].
16.
Martin, M,
Metzger DJ,
Michalek SM,
Connell TD,
and
Russell MW.
Comparative analysis of the mucosal adjuvanticity of the type II heat-liable enterotoxins LT-IIa and LT-llb.
Infect Immun
68:
281-287,
2000
17.
Moldoveanu, Z,
Vzorov AN,
Huang WQ,
Mestecky J,
and
Compans RW.
Induction of immune responses to SIV antigens by mucosally administered vaccines.
AIDS Res Hum Retroviruses
15:
1469-1476,
1999[ISI][Medline].
18.
Ng, WH,
Lutsar I,
Wubbel L,
Ghaffar F,
Jofri H,
McCracken GH,
and
Friedland IR.
Pharmacodynamics of trovafloxacin in a mouse model of cephalosporin-resistant Streptococcus pneumoniae pneumonia.
J Antimicrob Chemother
43:
811-816,
1999
19.
Simecka, JW,
Jackson RJ,
Kiyono H,
and
McGhee JR.
Mucosally induced immunoglobin and E-associated inflammation in the respiratory tract.
Infect Immun
68:
672-679,
2000
20.
Stampfli, MR,
Cwiartka M,
Gajewska BU,
Alvarez D,
Ritz S,
Inman MD,
Xing Z,
and
Jordana M.
Interleukin-10 gene transfer to the airway regulates allergic mucosal sensitization to mice.
Am J Respir Cell Mol Biol
21:
586-596,
1999
21.
Takafuji, S,
Suzuki S,
Koizumi K,
Tadokoro K,
Miyamoto T,
Ikemori R,
and
Muranka M.
Diesel exhaust particulates inoculated by the intranasal route have an adjuvant activity for IgE production in mice.
J Allergy Clin Immunol
79:
639-645,
1987[ISI][Medline].
22.
Tebbey, PW,
Unczar CA,
LaPierre NA,
and
Hancock GE.
A novel and effective intranasal immunization strategy for respiratory syncytial virus.
Viral Immunol
12:
41-45,
1999[ISI][Medline].
23.
Tsuyuki, S,
Tsuyuki J,
Einsle K,
Kopf M,
and
Coyle AJ.
Costimulation through B7-2 (CD86) is required for the induction of a lung mucosal T helper cell 2 (Th2) immune response and altered airway responsiveness.
J Exp Med
185:
1671-1679,
1997
24.
Wagner, EM,
and
Foster WM.
Interdependence of bronchial circulation and clearance of 99mTc-DTPA from the airway surface.
J Appl Physiol
90:
1275-1281,
2001
25.
Walther, S,
Wenyao S,
and
Lennquist S.
Pulmonary dynamics of radiolabelled erythrocytes and leucocytes in early gram-negative sepsis in pigs.
Eur J Surg
165:
979-985,
1999[ISI][Medline].
26.
Zhang, Y,
Lamm WJE,
Albert RK,
Chi EY,
Henderson WR,
and
Lewis DB.
Influence of the route of allergen administration and genetic background on the murine allergic pulmonary response.
Am J Respir Crit Care Med
155:
661-669,
1997[Abstract].