
==== Front
Animal Model Exp Med
Animal Model Exp Med
10.1002/(ISSN)2576-2095
AME2
Animal Models and Experimental Medicine
2096-5451
2576-2095
John Wiley and Sons Inc. Hoboken

38925626
10.1002/ame2.12460
AME212460
AMEM-2024-0053.R1
Original Article
Regular Article
Original Article
An ethically guided preclinical device for phenotyping H2 production in laboratory rodents
Pascal‐Moussellard et al.
Pascal‐Moussellard Victor 1
Boucher Emilie 1
Tanguy Stéphane 1
Cinquin Philippe 1
Barraud Pierre‐Alain 1
Davin Chloé 1
Salomez‐Ihl Cordélia 1
Hannani Dalil 1
Boucher François 1
Alcaraz Jean‐Pierre https://orcid.org/0000-0002-5924-9519
1 jean-pierre.alcaraz@univ-grenoble-alpes.fr

1 University Grenoble Alpes, CNRS, UMR 5525, VetAgro Sup, Grenoble INP, TIMC Grenoble France
* Correspondence
Jean‐Pierre Alcaraz, University Grenoble Alpes, CNRS, UMR 5525, VetAgro Sup, Grenoble INP, TIMC, Grenoble 38000, France.
Email: jean-pierre.alcaraz@univ-grenoble-alpes.fr

25 6 2024
8 2024
7 4 10.1002/ame2.v7.4 Themed Issue: Study on Cardiovascular and Cerebrovascular Diseases 553561
23 2 2024
03 6 2024
© 2024 The Author(s). Animal Models and Experimental Medicine published by John Wiley & Sons Australia, Ltd on behalf of The Chinese Association for Laboratory Animal Sciences.
https://creativecommons.org/licenses/by-nc-nd/4.0/ This is an open access article under the terms of the http://creativecommons.org/licenses/by-nc-nd/4.0/ License, which permits use and distribution in any medium, provided the original work is properly cited, the use is non‐commercial and no modifications or adaptations are made.

Abstract

Background

Dihydrogen (H2) is produced endogenously by the intestinal microbiota through the fermentation of diet carbohydrates. Over the past few years, numerous studies have demonstrated the significant therapeutic potential of H2 in various pathophysiological contexts, making the characterization of its production in laboratory species of major preclinical importance.

Methods

This study proposes an innovative solution to accurately monitor H2 production in free‐moving rodents while respecting animal welfare standards. The developed device consisted of a wire rodent cage placed inside an airtight chamber in which the air quality was maintained, and the H2 concentration was continuously analyzed. After the airtightness and efficiency of the systems used to control and maintain air quality in the chamber were checked, tests were carried out on rats and mice with different metabolic phenotypes, over 12 min to 1‐h experiments and repeatedly. H2 production rates (HPR) were obtained using an easy calculation algorithm based on a first‐order moving average.

Results

HPR in hyperphagic Zucker rats was found to be twice as high as in control Wistar rats, respectively, 2.64 and 1.27 nmol.s−1 per animal. In addition, the ingestion of inulin, a dietary fiber, stimulated H2 production in mice. HPRs were 0.46 nmol.s−1 for animals under control diet and 1.99 nmol.s−1 for animals under inulin diet.

Conclusions

The proposed device coupled with our algorithm enables fine analysis of the metabolic phenotype of laboratory rats or mice with regard to their endogenous H2 production.

An innovative solution to accurately monitor H2 production in free‐moving rodents while respecting animal welfare standards. The proposed device coupled with our algorithm enables fine analysis of the metabolic phenotype of rodents with regard to their endogenous H2 production rates.

hydrogen therapy
laboratory rodents
microbiota
molecular hydrogen
noninvasive monitoring device
Université Grenoble Alpes through the Cross Disciplinary Program (CDTools) “My Health Companions”. source-schema-version-number2.0
cover-dateAugust 2024
details-of-publishers-convertorConverter:WILEY_ML3GV2_TO_JATSPMC version:6.4.8 mode:remove_FC converted:03.09.2024
Pascal‐Moussellard V , Boucher E , Tanguy S , et al. An ethically guided preclinical device for phenotyping H2 production in laboratory rodents. Anim Models Exp Med. 2024;7 :553‐561. doi:10.1002/ame2.12460
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pmc1 INTRODUCTION

Dihydrogen (H2), more commonly called molecular hydrogen, is the most abundant molecule in the universe. On Earth, H2 is produced at high temperature from olivine during Earth's lithospheric mantle hydration at great depths and can serve as a potential source of energy for human activities. 1 , 2 Animals also produce H2. The intestinal microbiota of animals contains certain species of microorganisms that can produce H2 through hydrogenase‐mediated fermentation of carbohydrates. The activity of these microorganisms in the mammalian digestive tract is such that it is estimated that human microbiota is capable of producing over 1 L of H2 per 24 h. 3

The H2 produced in the intestine can have three outcomes. (1) It can be evacuated directly with intestinal gas. (2) It can be consumed by the energy metabolism of methanogenic bacteria. (3) It can cross the intestinal epithelium in dissolved form, diffuse into the internal environment, be distributed throughout the body, and finally be excreted mainly through the lungs. It can also be excreted, to a lesser extent, through the skin, 4 or in the urine. 5 In clinical practice, exhaled H2 has been used for more than 70 years as an easy way to diagnose carbohydrate malabsorption, for instance, lactose intolerance. 6 The use of the “breathing test” to assess exhaled H2 has more recently been extended to the diagnosis of intestinal dysbiosis, whose contribution to many chronic pathologies is the subject of a growing number of publications. 7 , 8 For these so‐called breathing tests, several tools have been developed and are available for clinical versions.

Over the past 15 years, numerous experimental and clinical studies have demonstrated (i) the link between microbiota and health, 9 , 10 (ii) the fact that H2 monitoring reflects the composition and activity of the gut microbiota, and (iii) the therapeutic potential of H2 in relation to its antioxidant, anti‐inflammatory, and antiapoptotic properties. 11 , 12 , 13 Toxicological and pharmacological studies have proven the safety of hydrogen, 14 and we have recently demonstrated its absence of genotoxicity in vivo in rats. 15 It is now becoming increasingly necessary to develop tools for measuring exhaled H2 in rodents to enable relevant preclinical approaches adapted to these emerging concepts. The main obstacle to the development of satisfactory hydrogen breath test tools for laboratory rodents is the impossibility of collecting exhaled gas samples from unanesthetized laboratory rodents. 16 A solution to the problem of collecting air samples in conscious animals was first proposed by Gumbmann and Williams. 17 This group pioneered a method for measuring exhaled H2 in rats using a noninvasive procedure for repetitive testing of the same animal under various nutritional interventions. The animal was placed in a bowl desiccator with an internal diameter of 20 cm. A 1‐mL air sample containing the accumulated H2 was collected every hour and analyzed using gas chromatography. Following this first study, several authors have proposed optimizations of the same setup, all based on timed collection, and not always fully satisfactory in terms of animal welfare. 18 , 19 , 20 , 21

Until now, none of the existing systems have combined ethical aspects of animal welfare and accurate real‐time monitoring of the flow of H2 exhaled by an animal. The aim of the present study was to develop and validate an original preclinical device that incorporates physiological and ethical features and enables continuous monitoring of exhaled H2 in laboratory rodents. For the validation stage, we chose rats and mice with different metabolic phenotypes likely to impact H2 content in exhaled air to challenge our device in terms of sensitivity and robustness.

2 METHODS

2.1 Device setup

The proposed H2 monitoring device is shown in Figure 1. It was composed of a gastight chamber (glass desiccator, total internal volume 9.4 L) in which a mesh‐cage of 8.1‐L internal volume was placed to receive the animals. Two standard 92‐mm fans were placed in the chamber to homogenize the air without disturbing the animals. To achieve this, the first fan was fitted with a support to direct the airflow laterally, and the second was placed under the cage to ensure downward air movement.

FIGURE 1 Global view of the H2 monitoring device: (1) H2 sensor ATO SKY 2000, (2) multisensor (O2, CO2, relative humidity and temperature), (3) cage, (4) upper fan, (5) lower fan coupled with the CO2 absorption system (soda lime), (6) H2O absorber (silica gel), (7) data collection, (8) power supply, (9) peristaltic pump, (10) O2 container, and (11) active charcoal filters.

CO2 and H2O are physiologically produced by cellular metabolism. Thus, when an animal is placed in a hermetic enclosure, CO2 accumulates rapidly, and the atmosphere becomes humid. CO2 is an asphyxiating and toxic gas, and humidity runs the risk of damaging the sensors of the device. To maintain CO2 within the atmospheric range—around 500 ppm—175‐g soda lime (Spherasorb Intersurgical) was placed onto a carbon filter, above the lower fan. H2O accumulation was prevented by adding 150‐g silica gel at the bottom of the chamber.

A gastight Tedlar bag containing 2‐L O2 (generated by a 525 KS Drive Devilbiss concentrator) was connected to a peristaltic pump, itself connected to the chamber with a 1/1.8‐mm polyethylene tubing, allowing the control of the supply of oxygen to the animals.

O2 and CO2 levels, as well as relative humidity and temperature, were constantly measured in the chamber using a homemade, optimized multisensor detector housed on the top of the cage. The sensors (DFROBOT: SEN0322 for O2, SEN0159 for CO2, and SEN0137 for relative humidity and temperature) were mounted on a board and managed by an open‐source microcontroller (Arduino). Data acquisition was handled by the Arduino and transmitted to the PC software via a USB port. This software acquired the data of the three sensors on an SD card at 1 Hz. During the whole experiment, O2 and CO2 concentration, temperature, and relative humidity were monitored.

Two H2 gas detectors, “SKY2000,” were purchased from ATO (USA). The “ATO 500” (range 0–500 ppm, detection threshold 1.6 ppm, accuracy 0.1 ppm) was used for rat measurements, and the “ATO 100” (range 0–100 ppm, detection threshold 0.25 ppm, accuracy 0.1 ppm) was used for mice. Before use, ATO H2 sensors were calibrated against a Quintron BreathTracker, which was used as a reference. The ATO H2 sensor was placed on a bypass, and its pump was set at 22.5 L.h−1 to ensure airflow within the bypass.

Two 50‐mL syringes containing an activated carbon filter were placed on the in‐ and outlines of the bypass to capture contaminating gases—mainly ammonia—and to dampen the transmission of pump noise inside the chamber.

All electrical wiring (power supply and data collection) and tubing (O2 input and H2 measurement bypass lines) were routed through a rubber stopper that hermetically sealed the chamber lid.

2.2 Animal experiments

2.2.1 Ethical issues

Animal welfare is a growing concern in the design and development of new preclinical devices. The experiments involving animals presented in this article are limited to noninvasive explorations performed on conscious animals. In addition, the animals included in this study were reused from previous protocols to meet the principles of the three Rs (replace, reduce, refine). All experiments were carried out in accordance with the European Communities Council Directive 2010/63/EU (Directive 2010/63/EU, 2010). Rats were reused after two procedures previously authorized by the French Ministry of Research under the references #36969–202 204 251 517 299 (male Wistar rats, 8 weeks old, average body weight = 258 ± 12 g, n = 6) and #38828‐2022091611584664 (male Zucker [fa/fa] rats, 8–10 weeks old, average body weight = 538 ± 38 g, n = 6). For experiments in mice, 41‐week‐old C57BL/6 female mice (n = 24, 30–35 g) were reused from an earlier study (#26300‐2020070111383424) in which 12 mice were supplemented with inulin Orafti GR (Beneo) in drinking water (72 g.L−1) made available ad libitum for 41 weeks, whereas the other 12 (control group) received regular water over the same period. 22

2.2.2 Monitoring of H2 concentration in the device

Rats were individually placed in the experimental setup at different times of the day, and H2 production was followed for 1 h. The two experimental groups of mice studied were subdivided into two batches of six mice each for H2 measurements. Each batch of six mice was measured for 12 min. The measurements were made twice a day, at 7 a.m. and 1 p.m.

2.3 Data analysis

A simple calculation algorithm was developed to determine the H2 production rate (HPR) from the continuous measurement of H2 concentration in the device (Figure 2). H2 production curves were first converted from parts per million to nanomoles of H2 as a function of time. The raw data expressed in parts per million were first multiplied by the volume of the enclosure (V e  = 9.4 L) to convert them into microliters. Considering H2 as a perfect gas in standard conditions of ambient temperature and pressure, dividing the volumes of H2 by 24.5 L (volume of 1 mol of gas) and multiplying by 1000 converted the data into nanomoles. The nanomole H2 dataset was then processed by performing a first‐order moving average (MA[1]) over a 10‐point sliding window. HPR (expressed in nmol.s−1) was calculated as the slope of the linear regression of the nanomole H2 dataset as a function of the time spent in the chamber, starting at the first non‐null point.

FIGURE 2 Data processing for H2 production rate (HPR) calculation. Raw dataset of H2 concentration ([H 2 ] n ) given by the sensor was first converted from parts per million into nanomoles considering 24.5 L as the volume of 1 mol of H2 under standard conditions of ambient temperature and pressure. The nanomole H2 dataset (X n ) was then processed by performing a first‐order moving average (MA[1]) over a 10‐point sliding window (W n ). HPR expressed in nmol.s−1 was calculated as the slope of the linear regression of the MA(1) nmol H2 dataset (Xn¯) as a function of the time spent in the chamber (t), starting from the first non‐null point. V e : enclosure volume (9.4 L).

2.4 Statistics

Paired t‐test was used to statistically compare the HPR of Wistar rats in the morning and in the afternoon. Independent t‐test was used to statistically compare the HPR of Zucker rats to that of Wistar rats.

3 RESULTS

3.1 Device characterization and air quality control

The gas tightness of the device (Figure 1) was investigated by monitoring the stability of H2 concentration after injecting 50, 100, or 200 ppm of H2 in the chamber (Figure 3A). After an initial rapid increase in H2 concentration at the time of injection and a slight transient decrease related to gas homogenization in the device, H2 reached a plateau.

FIGURE 3 H2 monitoring device characterization. (A) H2—leak test. Continuous recording after injection of 200 (black circles), 100 (empty circles), or 50 ppm of H2 (empty triangles) in the hermetic chamber. Arrows correspond to the start of the steady state. (B) CO2 measurements of a rat into the chamber in absence of soda lime. (C) Absorption of CO2 of human exhaled air into the chamber in the presence of 200‐mL soda lime. (D) Environmental parameters in the chamber. O2 (empty triangles), CO2 (black circles), humidity (back squares), and temperature (empty diamonds) were measured for 1 h in the presence of six C57BL/6 mice.

The CO2 production of a rat of 260‐g body weight was measured for 2 min with the CO2 detector in the chamber from which the soda lime had been removed. The rat produced 3.1 μmol.s−1 of CO2 (Figure 3B). The CO2 absorption capacity of the CO2 absorption system was tested by increasing CO2 concentration up to >6000 ppm in the chamber. The observed maximal absorption rate of CO2 in the device was 18.4 μmol.s−1 (Figure 3C).

Air composition in the chamber was monitored with mice (Figure 3D). The O2 content in the chamber was manually maintained between 19% and 23% by adapting the flow rate of the peristaltic pump to the instantaneous O2 value. Experiments confirmed the efficiency of the CO2 absorption system as the CO2 concentration was maintained around the normal atmospheric concentration of 500 ppm (plateau value: 490 ± 17 ppm; Figure 3D). Finally, relative humidity was maintained below 80%, thanks to silica gel, and the temperature remained constant throughout experiments (Figure 3D).

3.2 Increased HPR in hyperphagic rats

In both control and hyperphagic rats, H2 production was found to be almost stationary during the hour of investigation (Figure 4A). This enabled us to determine the HPR as described in Section 2.3.

FIGURE 4 H2 production rate (HPR) monitoring in rats. (A) The raw data of H2 production of one Zucker (black circles) and one Wistar (empty circles) rat measured during 1 h. (B) HPR of Wistar rats, measured in the morning (empty column, n = 6) or in the afternoon (hatched column, n = 6), and that of Zucker rats (black column, n = 6). Paired and independent t‐test were performed, *p‐value < 0.05, **p‐value < 0.01. Data are represented as mean ± standard deviation (SD).

No significant difference in mean HPR was observed in control Wistar rats between morning, 1.25 ± 0.71 nmol.s−1, and afternoon measurements, 1.44 ± 1.00 nmol.s−1 (Figure 4B). However, the HPR in hyperphagic Zucker rats, 2.58 ± 0.72 nmol.s−1, was, on average, twice that of age‐matched control Wistar rats, whether they were measured in the morning or in the afternoon, p < 0.01 (Figure 4B).

3.3 Inulin supplementation induces a cyclic increase in HPR over the nychthemeral period in mice

In the mouse study, HPR was measured on individual animals as well as on batches of six animals. Both approaches enabled HPR to be determined provided the measurement time was sufficient. Our calculation algorithm proved to be robust enough to determine the very low HPR of a single control C57BL/6 mouse placed in the device for 1 h (HPR = 5.60 10−2 nmol.s−1; Figure 5A).

FIGURE 5 Monitoring of H2 production rate (HPR) in mice. (A) First‐order moving average of H2 production of a single C57BL/6 control mouse. Data processing is described in the Materials and Methods section. (B) Raw data of H2 production of two batches of six control C57BL/6 mice (empty circles and squares) or two batches of six C57BL/6 mice supplemented with 72 g.L−1 inulin ad libitum (black circles and squares) between 7 a.m. and 8:30 a.m. (C) Raw data of H2 production of two batches of six control C57BL/6 mice (empty circles and squares) or two batches of six C57BL/6 mice supplemented with 72 g.L−1 inulin ad libitum (black circles and squares) between 1 p.m. and 2:30 p.m.

For measurements carried out on batches of mice, a wide variation in HPR as a function of time of day was observed for animals chronically treated with inulin for 41 weeks, whereas this daily variation was minimal in age‐matched control mice. Thus, overall HPR (for a batch of six mice) measured at 7 a.m. was 8.7 ± 3.4 nmol.s−1 in the two batches of inulin‐supplemented mice (i.e., normalized HPR per mouse = 1.45 ± 0.57 nmol.s−1) and 2.8 ± 0.9 nmol.s−1 in the two control batches (i.e., normalized HPR per mouse = 0.47 ± 0.15 nmol.s−1; Figure 5B). In contrast, overall HPR at 1 p.m. was 1.5 ± 0.32 nmol.s−1 for the two inulin batches (i.e., normalized HPR per mouse = 0.25 ± 0.05 nmol.s−1) and 1.5 ± 0.1 nmol.s−1 for the two control batches (i.e., normalized HPR per mouse = 0.25 ± 0.02 nmol.s−1; Figure 5C).

To complete the nutritional approach in mice, inulin was withdrawn for 24 h (Figure 6A) in both batches treated for 41 weeks. The HPR calculated for mice in the inulin and control batches was very similar regardless of the time of day (overall HPR in the four batches: 2.0 ± 0.5 nmol.s−1 at 7 a.m., that is, normalized HPR per mouse = 0.33 ± 0.08 nmol.s−1, and 2.0 ± 0.4 nmol.s−1 at 1 p.m., that is, normalized HPR per mouse = 0.33 ± 0.07 nmol.s−1). The drinking water of all experimental groups (control and inulin) was then supplemented with inulin (72 g.L−1) for the following 24 h. HPR in the two combined groups of mice was 11.7 ± 3.2 nmol.s−1 at 7 a.m. (i.e., normalized HPR per mouse = 1.95 ± 0.53 nmol.s−1) and 1.6 ± 0.4 nmol.s−1 at 1 p.m. (i.e., normalized HPR per mouse = 0.27 ± 0.07 nmol.s−1) (Figure 6B).

FIGURE 6 Inulin‐induced H2 production monitoring. (A) Mice fed a control diet (control 1 and 2, empty circles and squares) or supplemented with inulin (inulin 1 and 2, black circles and squares) for 41 weeks received a control diet for 24 h during which H2 production was measured over 12 min, once between 7 a.m. and 8:30 a.m. and once between 1 p.m. and 2:30 p.m. (B) Mice fed a control diet (control 1 and 2, empty circles and squares) or supplemented with inulin (inulin 1 and 2, black circles and squares) for 41 weeks received a control diet for 24 h and were then supplemented with inulin (72 g.L−1) for the next 24 h. During the 24‐h inulin supplementation, H2 production was measured over 12 min, once between 7 a.m. and 8:30 a.m. and once between 1 p.m. and 2:30 p.m.

4 DISCUSSION

The preclinical device developed in this work meets the objectives initially defined. First, it gives access to a new physiological variable, the HPR, and proposes a calculation algorithm enabling its robust determination. Second, the proposed new system allows physiological measurements to be carried out under satisfactory ethical conditions, guaranteeing both the animals' physical integrity and well‐being. Furthermore, the experimental validation study proposed in this work was carried out on animals from previous protocols to respect the three Rs rule. Finally, we provide examples of preclinical situations in which our device was successfully used to characterize rodents with different experimental phenotypes by measuring their HPR.

All previous attempts to measure H2 production in preclinical studies were based on timed collection methods with either a very limited number of air samples or even a single one. 18 , 19 , 20 , 21 Our device allows a continuous measurement of H2 accumulation in the airtight enclosure at an acquisition frequency of 0.2 Hz. From the raw data produced by the sensor, a simple calculation algorithm delivers a HPR value corresponding to the slope of the first‐order moving averages of H2 accumulation over time. This method increases both the robustness and accuracy of the measurement.

The device consists of a hermetically sealed enclosure in which the animal(s) is/are confined for the duration of the acquisition. The environmental variables in which the animals are placed are perfectly controlled throughout the measurement period, in particular the composition of the air in terms of oxygen, CO2, and other contaminant gases, temperature, pressure, hygrometry, and noise level.

To demonstrate the suitability of our device under conditions of use, we first carried out HPR measurements in a rat model of hyperphagia. Our preclinical device evidenced that HPR of Zucker rats is twice as high as that of lean control Wistar rats of the same age. Zucker rats carry a mutation in the gene encoding the receptor of leptin, which makes them hyperphagic. At 10 weeks of age, male Zucker rats are obese and diabetic, 23 and their daily food intake is 30 g a day, almost 35% more than age‐matched lean rats. 24 In addition, the microbiota of Zucker rats shows some particularities, including an enrichment in Bacteroidota and Bacillota bacterial phyla compared to lean rats. 25 Surprisingly, the increase in HPR (50%) measured in Zucker rats was greater than the described increase in food intake (35%). We propose that this lack of proportionality between the two phenomena is due to the qualitative changes in the colonic microbiota characteristic of the Zucker rat phenotype, because Bacteroidota and Bacillota, the favored bacterial phyla, are highly H2‐producing. 26 Further studies will be required to verify this hypothesis. Nevertheless, this study shows that our system can accurately assess the HPR of a single rat over a 1‐h measurement. Moreover, by placing a single mouse in our device, we further demonstrated that it was also possible to quantify very low values of HPR within an hour, thanks to the calculation algorithm we developed. Indeed, if we consider the accuracy (0.1 ppm) and detection threshold (0.25 ppm) of the ATO 100 sensor, we can estimate the sensitivity limit of our device using our algorithm over a 1‐h measurement, at 4 10−2 nmol.s−1 (0.1 + 0.25 = 0.35 ppm.h−1 = 10−4 ppm.s−1 = 4 10−2 nmol.s−1). Single‐animal measurement paves the way to accurate and noninvasive phenotyping of laboratory animals in studies on the microbiota.

Under certain conditions of use, phenotyping batches of six mice may offer an advantage over single‐animal measurements, as it enables an average HPR value to be obtained in a single rapid measurement lasting no more than 12 min. Using this method, we were able to determine a normalized HPR as low as 0.2 nmol.s−1 per mouse, a value unattainable with existing devices. Measuring the overall HPR on a batch of mice can therefore be a considerable asset, for example, to determine the effects of a nutritional intervention or pharmacological treatment rapidly, without reducing the quality of the physiological measurement.

To further demonstrate the efficacy and flexibility of our device, we chose to implement a nutritional manipulation on mice. In this particular context, the proposed preclinical device revealed that H2 production is stimulated for a specific time period following inulin intake. Inulin is a dietary fiber mainly found in vegetable roots and exhibits prebiotic activity. Its hydrolysis by colonic flora releases fructose, which is thereafter used by certain H2‐producing microbial species. Inulin can be used clinically for H2 breath test and for the diagnosis of fructan intolerance. 27 , 28 Our group and others have already demonstrated that, in addition to stimulating H2 production in the gut, 29 an inulin‐enriched diet has the ability to modulate the composition of the gut microbiota. 22 , 30 , 31 The process of inulin digestion in mice has recently been investigated using fluorescent markers, demonstrating that its colonic digestion phase begins 2 h after the ingestion and peaks after 4.5 h. 32 It can therefore be anticipated that a peak H2 production should occur 4.5 h after inulin ingestion. As mice are semi‐nocturnal rodents, they eat and drink mainly at night. For this reason, H2 production by mice supplemented with inulin in their drinking water should peak in the early morning and fall to a minimum at the end of the day. Our results show that, as expected, at 7 a.m., inulin‐supplemented mice had a normalized HPR per mouse more than three times higher than mice fed a control diet. Again, as expected, at 1 p.m., that is, at a distance from food and drink intake, normalized HPR per mouse in inulin‐supplemented mice was low and close to that measured in control mice. As inulin promotes the growth of specific bacteria in the gut microbiota, 22 , 30 , 31 we could speculate that the difference in HPR between mice chronically treated with inulin and control mice could mainly stem from the composition of the microbiota. To address this issue, inulin chronic supplementation was interrupted for 24 h prior to HPR measurement. Interestingly, both experimental groups showed equivalent and low levels of HPR. Immediately after the HPR measurement, both experimental groups received inulin‐enriched drinking water for the following 24 h. This second phase of the experiment corresponded to a resumption of chronic treatment for mice of the inulin‐supplemented group and acute treatment for mice of the control group. HPR, assessed at 7 a.m., was identical in the two groups and five times higher than the basal value measured the previous day. Moreover, the HPR value obtained at this stage of the experiment was equivalent to that obtained in the inulin‐supplemented group just prior to treatment interruption. When HPR was measured at 1 p.m., it was significantly decreased in both groups to basal levels, as previously observed in the inulin‐supplemented group.

Altogether the measurements we have carried out under operational preclinical conditions show that the proposed device can acquire accurate data in different rodent species and under various physiological conditions. Its high sensitivity enables accurate assessment of the H2 production rate of small laboratory animals, even at very low levels. Finally, our device enables direct HPR determination of single and grouped animals over a short measurement period.

5 CONCLUSIONS

This study demonstrates the efficacy, robustness, and sensitivity of our device, which is capable of monitoring HPR in laboratory rodents under a variety of preclinical conditions. Our validation study demonstrates the relevance of HPR as a first exploratory approach to gut microbiota phenotype in nutritional models and in animals with particular genotypes. Implementing additional sensors to monitor other exhaled gases could further enhance the relevance of the device in the context of completely noninvasive phenotyping of laboratory animals. The use of the proposed device could also be extended to the development and characterization of new hydrogen therapy treatments or devices, a currently promising sector, in particular, for the treatment of inflammatory pathologies or those with a strong oxidative component.

AUTHOR CONTRIBUTIONS

Conceptualization and methodology: Victor Pascal‐Moussellard, Emilie Boucher, Stéphane Tanguy, Philippe Cinquin, Pierre‐Alain Barraud, Chloé Davin, Cordélia Salomez‐Ihl, François Boucher, and Jean‐Pierre Alcaraz. Investigation and formal analysis: Victor Pascal‐Moussellard, Emilie Boucher, Stéphane Tanguy, François Boucher, and Jean‐Pierre Alcaraz. Writing—original draft preparation: Victor Pascal‐Moussellard and Jean‐Pierre Alcaraz. Writing—review and editing: Victor Pascal‐Moussellard, Emilie Boucher, Stéphane Tanguy, Philippe Cinquin, Pierre‐Alain Barraud, Chloé Davin, Cordélia Salomez‐Ihl, Dalil Hannani, François Boucher, and Jean‐Pierre Alcaraz. Project administration: Philippe Cinquin and François Boucher. Funding acquisition: Philippe Cinquin. All authors have read and agreed to the published version of the manuscript.

FUNDING INFORMATION

This research was funded by Université Grenoble Alpes through the Cross Disciplinary Program (CDTools) “My Health Companions”.

CONFLICT OF INTEREST STATEMENT

The authors declare no conflict of interest.

ETHICS STATEMENT

The study was conducted in accordance with the Declaration of Helsinki and approved by the Ethics Committee #12 of Grenoble Alpes University and by the French Ministry of Higher Education and Research.

ACKNOWLEDGMENTS

The authors thank the Laboratoire Radiopharmaceutiques Biocliniques (LRB) for supplying the Zucker rats and the corresponding Wistar lean controls. Orafti GR inulin was generously donated by Beneo GmbH.

DATA AVAILABILITY STATEMENT

The datasets obtained and/or analyzed during the current study are available from the corresponding author on reasonable request.
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