Plethysmograph training: A refinement for collection of respiration data in mice

Abstract

Inhaled chemicals can harm the airways. Different effects can result in distinct changes in respiratory patterns; the type of change indicates where and how the respiratory system is affected. Furthermore, changes in respiratory patterns may be detected at much lower substance concentrations than those that cause more serious effects, such as histopathological changes. Changes in respiratory patterns can be studied experimentally by monitoring the breathing of mice placed in plethysmographs and exposing head-out to the test substance. The method is well established; however, it is not known if training mice in being restrained in the plethysmograph could increase the quality of data collection. Here we report the results of training mice to be restrained in plethysmographs for 5 consecutive days, with respect to body weight, respiratory parameters, and time spent in the plethysmograph, before they are removed because of unstable breathing patterns. The mice tolerated the procedure better (measured by time in the plethysmograph) on the second day of training than the first day. Training did not change the breathing parameters between days. Breathing parameters stabilized within 5 min after the mice were placed in the plethysmographs on all days. There was an average of 3% weight loss between the first and last days of the training, indicating that the training procedure placed some strain on the animals. Training reduces the number of mice attempting to escape from the plethysmograph.

INTRODUCTION

Restraining mice in whole-body plethysmographs is a method used for measuring changes in respiration in conscious and freely breathing animals. The method is highly useful when studying the effects of different substances on the respiratory system, as distinct changes in different respiratory parameters occur depending on which area of the respiratory system is affected.^1,2

Sensory irritation is caused by the substance interacting with the trigeminal nerve endings in the eyes and upper respiratory tract. Sensory irritation is one of the most frequently used endpoints for setting occupational exposure limits (OELs), accounting for an estimated 40% of all OELs; the data on irritation are from human reports as well as from animal data.³ In humans, sensory irritants cause a painful and burning sensation in the eyes, nose, and throat as well as coughing, depending on the concentration of the irritant. The American Society for Testing and Materials (ASTM) International Standard number E981-19 “Standard Test Method for Estimating Sensory Irritancy of Airborne Chemicals”⁴ describes how changes in respiration in mice can be used to assess sensory irritation. The method determines the potency of the irritant by determining the aerosol exposure concentration that results in a 50% reduction in repsiratory rate (RD₅₀) for a substance.⁵ There is a close correlation between the RD₅₀ values measured in exposed mice and the American threshold limit values for the same substance^6,7; they have been used to set OELs⁶ and have been correlated with no observed adverse effect levels in humans to determine acceptable air exposure of the general public.⁸ Measuring changes in breathing patterns in the Alarie assay can also indicate if the exposure has pulmonary irritating effects or if it limits airflow.¹

The Organization for Economic Co-operation and Development regulatory test guidelines (TG) are used to investigate hazardous properties of chemicals, and the results are used for classifying and labeling among other purposes. Experimental animals exposed to substances via inhalation using regulatory accepted guidelines for acute inhalation toxicity^9–11 are observed during exposure and 14 days postexposure, and the endpoint is death,^9,10 or evident signs of toxicity (TG433¹¹). For TG433 the most predictive evident signs of toxicity are cage-side observation of irregular respiration, hypoactivity, tremors, and body weight loss >10%.^12,13 Changes in respiratory patterns often occur at much lower aerosol concentrations than concentrations that cause more serious effects, for example, for plant protection products.¹⁴ Therefore, about 90% of the compounds tested for inhalation toxicity induced local effects in the lungs at lower concentrations than those causing 50% lethality.¹⁵

The initial phase of irritation is stimulation of the nerve endings in the respiratory tract and the eyes, for example, triggering cough and eye blinking; the second phase is tissue irritation, where the cells are affected. Neural activation is not in itself adverse, but if it is prolonged, it can impede reversibility³; both nerves in the upper respiratory tract and eyes and in the lower respiratory tract can be stimulated by inhaled substances depending on where in the system they deposit.⁶ Currently, the regulation for the classification, labeling, and packaging of substances and mixtures do not recognize the ASTM standard⁴ as a measure for irritation as the regulation relies only on human data, or cytotoxicity of the airways (section 3.8.2.2.1 “Criteria for respiratory tract irritation”¹⁶).

Monitoring breathing patterns using plethysmographs to assess the acute effects of substances on respiration has been used previously, for example, for indoor air quality, impregnation products, nanoparticles, and pharmaceutical enhancers.^1,17–22 Some impregnation products caused a sudden and irreversible decrease in tidal volume, which was followed by death of the animals unless the exposure was terminated immediately. This was later found to be triggered by inhibition of lung surfactant function.¹⁸

The standardized method for determining RD₅₀⁴ stipulates that the mice acclimate in the plethysmograph for 10 min before starting the test. We have used plethysmographs extensively and have observed that very often at least one animal does not acclimate to the plethysmograph. These mice are removed during the baseline period because stable breathing patterns cannot be obtained. This causes delays in the experiments, for example, if new mice have to be included in the experiment, thus restarting the baseline period. Restraint is a potent stressor in rodents, but the response to the stress generally decreases with repeated exposure to the same stressor.^23,24 We hypothesized that the method could be refined by training the mice in the procedure (being restrained in the plethysmograph) before starting the actual exposure experiment. Therefore, we trained all mice that were to be included in a subsequent inhalation experiment, to evaluate the effect of training on acclimating to the procedure and to determine how many days of training would be needed to refine the method.

MATERIALS AND METHODS Animals

A total of 37 inbred BALB/cJRj male mice aged 6–7 weeks at arrival were purchased from Janvier, and they were housed in clear 1290D Euro standard type 3 cages furnished with aspen bedding, enriched with small aspen blocks (both from Tapvei) and nesting material (Enviro Dri, Lillico, Biotechnology). The photoperiod was from 6:00 a.m. to 6:00 p.m., the temperature was 21.6 ± 0.5°C, and the relative humidity in the room was 50 ± 5% (mean ± standard deviation). The cages were sanitized twice weekly. Food (Altromin no.: 1324, Altromin) and municipal tap water were available ad libitum. The mice were randomly assigned to cages on arrival, two to three mice per cage, and acclimatized for a minimum of 1 week before training in the plethysmographs. The training was performed using a staggered start, to coincide with exposures in the main experiment (this has been published²⁵). Mice were marked on the tail with a waterproof marker to ensure identification throughout the experiment. The training in the plethysmographs occurred between 8:00 a.m. and 12:00 p.m. The weight of the mice on day 1 of training was 24.3 ± 1.6 g.

Ethical statement

Animal experiments were performed in accordance with procedures approved by the Animal Experiments Inspectorate, Denmark (permission no.: 2019-15-0201-00114). All experiments were performed by trained personnel and conformed to the Danish Regulations on Animal Experiments (LBK no.: 474 af 15/05/2014 and BEK no.: 12 af 07/01/2016) as well as the EU directive 2010/63 on the protection of animals used for scientific purposes,²⁶ which include guidelines for care and use of animals in research. The animal experiments were approved by the local animal ethics committee before the animals were purchased. Anesthesia was not used during the experiments, because the bioassay requires the animals to be fully awake with uncompromised breathing.

During plethysmograph training, the mice were removed from the plethysmograph and returned to their cage if the tidal volume decreased suddenly and did not normalize after the position of the animal was adjusted in the plethysmograph, or if the mouse tried to escape by pulling its head into the plethysmograph and the breathing parameters did not normalize after the head was adjusted. If the mouse tried to escape into the exposure chamber, it would be killed immediately as the constrictive pressure on the chest causes terminal damage.

Plethysmograph training

The subsequent inhalation exposure experiment is described elsewhere²⁵; the exposure setup is shown in Figure 1. Before the main experiment, the mice were trained in the procedure of being restrained in plethysmographs. In the main experiment the mice were exposed 30 min per day; we chose to train the animals for 30 min each day to match the time of exposure. The exposure experiment had a staggered start; therefore, the training was also staggered, so that the interval between the last training day and the start of the exposure experiment was a maximum of 1–2 days.

FIGURE 1. A figure showing the exposure setup.

Thirty-seven mice were trained on five consecutive workdays as follows. The mouse was weighed and placed in a whole-body plethysmograph with an appropriate–sized neckhole. The plethysmograph is a glass tube with a tapered end, at which a rubber gasket with a hole is secured. The head of the mouse goes through the hole, and the gasket makes an airtight seal for the body of the mouse, restraining the mouse in a “head-out” position. The plethysmograph was placed in the exposure chamber head-out, and the mice were exposed to clean air flowing through the exposure device to be used in the main experiment, to generate the same level of noise. The lighting, airflow, and temperature were the same as for the subsequent exposure experiment. A timer was started as soon as all mice (n = 7–8) were placed in the exposure chamber. The mice were exposed to air for 30 min before they were removed from the plethysmograph and returned to cages. During the 30-min period, the breathing pattern was continuously monitored using the computer program Notocord Hem (version 4.4.0.2, Notocord Systems SA). The average of each parameter was calculated for each minute of measure. Mice that had unstable breathing patterns, spontaneously or from escape behavior, even after adjusting the head position in the plethysmograph, were removed prematurely, and the time for removal was noted. Mice that were removed prematurely participated in the training the next day. The respiration files for all mice were analyzed for respiratory rate (BPM), tidal volume (VT), time of break (TB), time of pause (TP), time of expiration (TE), time of inspiration (TI), and bronchoconstriction (VD).

RESULTS

The mice were weighed daily during the training period, and the average loss of weight from days 1 to 5 was 2.7 ± 3.5%. The average change in weight on consecutive days was −1 ± 2.7%, indicating that some strain was associated with the daily handling, restraint, and exposure, even if the animals were exposed to air only. There was no clear trend in the weight changes as all mice both lost and gained weight and in no particular pattern from one exposure day to the next.

The training was planned so that each mouse should spend 30 min in the exposure tube (“time in the plethysmograph”). However, on day 1 the mice spent on average 24 min in the plethysmograph (range: 7–30 min) (Figure 2), and 35% of the mice had to be removed from the plethysmograph prematurely. The mice were removed if the breathing pattern was unstable even after adjusting the head and body position or if the pattern became unstable because the mouse attempted to escape. The mice mainly escaped by pulling their head into the plethysmograph or by attempting to escape into the chamber itself. The latter occurred only once during the total training time (i.e., once in 92.5 h that mice spent training in the plethysmographs). On days 2, 3, 4, and 5, the mice spent on average 29, 29, 30, and 28 min, respectively, in the plethysmograph, and 3%, 5%, 0% and 19% of the mice had to be removed prematurely.

FIGURE 2. Time spent in the plethysmograph. Each dot represents one mouse on a given day. The open diamond represents the mean time of all mice on the given day spent in the plethysmograph.

The respiratory rate decreased during the first 5 min of training on all days (Figure 3A). The standard deviation of the measurements overlapped on all 5 days; however, the average respiratory rate generally increases with consecutive days. The tidal volume increased within the first few minutes on most days (Figure 3B) but stayed more or less stable after this. The TB decreased during the first 10 min of exposure on all days (Figure 3C) and was stable for the remainder of the training. The TP was stable throughout the exposure on all days (Figure 3D). The peaks in TP on days 1 and 2 correspond to escape attempts and are therefore driven by a few mice, and the standard deviation of the data becomes correspondingly large. Bronchoconstriction is calculated as the flow rate at half the tidal volume during exhalation. It decreases during the first 10 min of exposure and then stabilizes (Figure 3E). The flow rate was the lowest on day 1 and subsequently increased with more training days. For all the respiratory parameters, the standard deviation for all minutes measured overlapped between training days. The largest difference in the mean values can be seen in the parameter “time of pause,” as this parameter becomes very elongated as the mouse tries to escape the restraint of the plethysmograph.

FIGURE 3. Breathing parameters. The mean and standard deviation of all mice for each breathing parameter on each training day: (A) respiratory rate, (B) tidal volume, (C) time of break, (D) time of pause, and (E) bronchoconstriction measured as respiratory flow rate. Day 1 is represented by green dots and lines, day 2 is represented by orange triangles, day 3 is represented by purple squares, day 4 is represented by pink crosses, and day 5 is represented by open light-green squares. The figure can be seen without standard deviation in Figure S1.

DISCUSSION

We hypothesized that the method could be refined by training the mice to stay in the plethysmograph before initiation of the actual exposure experiment. Therefore, all mice, to be included in a subsequent inhalation experiment, were trained to evaluate whether training was feasible and to determine the number of days of training required to refine the method. We trained the mice for 5 consecutive days and monitored body weight, time in the plethysmograph, and breathing parameters. On the first training day, the mice spent 24 min in the plethysmograph on average, and 35% of the mice were removed prematurely. Already by day 2, the mice spent nearly 30 min in the plethysmograph, and only 3% of the mice had to be removed before the 30 min. The time spent in the plethysmograph increased on the consecutive days, but on the fifth day of training, 19% of the animals had to be removed before 30 min. This indicates that by the fifth day of training the improvement in time spent in the plethysmograph was beginning to reverse. This is in line with the observation that desensitization to potent stressors, like restraint, may be achieved even with a single exposure.^23,24 We observed a mean weight loss of 3% between the start and end of training, whereas the day-to-day weight loss change was 0%–1%.

The results indicate that mice need to be trained once for 30 min in being restrained in the plethysmograph with air exposure, with the last day of training being no more than 2 days before the actual experiment. This improves the experiment, as the mice do not have to be removed from the experiment for reasons unrelated to aerosol exposure—something that can otherwise delay the start of the experiment, as mice have to be replaced, causing doubts about the effect of the test substance due to high variation in breathing parameters. The choice of a single day of training was based on the decrease in animals needing to be removed from 35% to 3% already after 1 day of training, with no additional improvement occurring during the subsequent days. Furthermore, a limited training period would cause less strain, determined from body weight data.

Several respiratory parameters changed during the first 5–10 min (Figure 3) as the animals became used to being restrained in the plethysmograph; however, according to the standard procedure the mice should acclimate to the plethysmograph for 10 min before the experiment. The current measurements show that during these 10 min the respiration parameters stabilize unless the animals attempt to escape. This confirms that a 10-min period in the standard procedure is relevant and adequate.

CONCLUSION

Training mice to be restrained in the plethysmograph in the exposure chamber under realistic exposure conditions, before exposure, once for 30 min, improved the results of this assay as it reduced the number of mice attempting to escape from the plethysmograph during the exposure experiment.

AUTHOR CONTRIBUTIONS

Jorid B. Sørli: conceptualization, methodology, validation, formal analysis, investigation, resources, data curation, writing—original draft, writing—review and editing, visualization, and funding acquisition. Karin S. Hougaard: methodology and writing—review and editing. Niels Hadrup: validation, formal analysis, resources, writing—review and editing, and funding acquisition.

ACKNOWLEDGMENTS

We thank Eva Terrida and Michael Guldbrandsen for technical assistance.

FUNDING INFORMATION

This work was supported by a grant from the Danish Working Environment Research Fund (project name Sikker-Motor; number: 29-2019-09) and by “FFIKA, Focused Research Effort on Chemicals in the Working Environment” from the Danish Government.

CONFLICT OF INTEREST STATEMENT

The authors declare no conflicts of interest.