Design and Validation of an Arduino-Based, Cost-Effective Heated Anesthesia Induction Chamber for Mice

Abstract

Rodents are highly susceptible to peri-anesthetic hypothermia because of their small body size, large surface area to volume/mass ratio, and anesthetic-induced thermoregulatory depression. Early heat loss during induction is rapid and can be difficult to reverse, emphasizing the need for simple, reliable warming solutions during anesthetic induction. We designed and validated a low-cost, Arduino-controlled heated induction chamber for mice and evaluated its safety and efficacy in preventing early hypothermia. The device uses proportional integral derivative control to regulate a heated chamber floor. Bench testing quantified warm-up time, steady-state stability, and the temperature offset between the embedded thermistor and surface temperature. In a randomized, paired, repeated-measures design, 10 adult C57BL/6 mice underwent 2 experimental conditions, unheated induction compared with heated induction, followed by a standardized 10-minute anesthetic maintenance phase on a prewarmed, heated surgical monitoring platform. Core body temperature was measured with intraperitoneal radio frequency identification transponders, while rectal temperature and physiologic parameters (peripheral oxygen saturation, heart rate, respiratory rate) were recorded during maintenance. Recovery time was measured from isoflurane discontinuation until the first voluntary movement. The heated chamber reached thermal stability within 10 minutes; at steady state, a constant average surface temperature near 37.5 °C was reliably obtained. During induction, unheated controls showed a rapid decline in core temperature, while during the heated-chamber induction, they maintained normothermia. The control group did not regain baseline temperature until several minutes into the maintenance phase and stabilized at values ∼1 °C lower than those of the heated group. Both groups remained physiologically stable during anesthetic maintenance and recovered within comparable times. This cost-effective, Arduino-based chamber prevented early hypothermia without compromising physiologic stability and represents a practical refinement to improve rodent anesthetic procedures, animal welfare, and experimental reproducibility.

Introduction

Rodents are uniquely vulnerable to hypothermia during anesthesia because of their small body size, high surface area to volume/mass ratio, and limited capacity for endogenous heat production. These characteristics promote rapid thermal exchange with the environment, causing core body temperature to fall much faster than in larger mammals. Even short anesthetic procedures can result in clinically significant hypothermia unless active warming is applied.^1,2 Although active warming strategies have improved intra-anesthetic temperature control, maintaining thermal stability from the onset of anesthesia remains a challenge in the field.

Heat loss in anesthetized rodents follows the same fundamental physical and thermodynamic principles described in human and veterinary medicine (radiation, convection, evaporation, and conduction), but in these species, the rate and magnitude of these pathways are greatly amplified by species-specific physiology and common anesthetic practices.^3,4 Conduction, the direct transfer of heat to cooler surfaces via physical contact, occurs rapidly in rodents because of their low thermal mass relative to surface area. Radiation, the loss of heat through the emission of infrared energy to cooler surroundings, contributes proportionally more to hypothermia in rodents due to their relatively large surface area.⁵ Convection, the transfer of heat through moving air or gas, plays a particularly important role during inhaled anesthesia induction, when a high fresh gas flow of cold, compressed oxygen mixed with anesthetic agents displaces the warm boundary layer of air around the body.⁶ Evaporation, the loss of heat as a liquid converts to vapor, is especially relevant during surgical anesthetic procedures, during surgical site preparation after fur removal and the application of surgical site disinfecting scrubs.^5,7,8

In addition, the anesthetic agents may amplify these physical heat losses by depressing hypothalamic thermoregulatory control, reducing metabolic heat production, and abolishing behavioral defenses such as huddling or seeking warmer surfaces.^9,10 Many agents also cause peripheral vasodilation, redistributing warm core blood to the skin and extremities where heat is lost most rapidly. This redistribution phase can account for up to 80% of total heat loss within the first hour of anesthesia.^6,11 Once established, hypothermia can be difficult to reverse and can confound physiologic readouts and welfare assessments.^10,12

The preoperative period, particularly the anesthesia induction phase, is a critical yet often unprotected window for heat loss. Within seconds of anesthetic induction onset, thermoregulatory defenses collapse, allowing all heat-loss mechanisms to act simultaneously while the redistribution rapidly transfers core heat to the periphery. Empirical data highlight this vulnerability: in C57BL/6 and nude mice undergoing isoflurane induction without warming, core temperature dropped by ∼2.8 °C within just 3 minutes.¹ In rats, anesthesia without thermal support led to a 4-10 °C temperature fall within the first 15-20 minutes.⁶ This decline is often exacerbated by preparatory tasks, including fur removal, antiseptic application, and placement onto unheated surfaces, commonly occurring before warming support is initiated, and resulting in hypothermic plateaus reached before surgery even begins.^2,5,7,8,13 Compounding this issue, warm-water blanket systems, commonly used for rodent anesthetic thermal support, show design-dependent variability and slow response times, limiting their effectiveness during the crucial early warming phase.¹¹

Several strategies have been explored to mitigate this early hypothermic shift. Prewarming, involving elevation of both peripheral and core temperatures before induction, can reduce the core-to-periphery gradient and delay hypothermia onset. In rats, it has been shown to prevent hypothermia and shorten recovery times, and even modest prewarming of 0.3 to 0.5 °C successfully delayed hypothermia, while prewarming to 40 °C extended threshold time from ∼7 to 19 minutes.^6,9

More recently, active warming during induction has been explored. Rather than increasing stored body heat beforehand, this approach minimizes losses during redistribution by providing warmed contact surfaces at the onset of anesthesia. Heated induction chambers or heating pads beneath the chamber have helped to maintain normothermia during short inductions, with unheated controls dropping ∼3 °C.¹ In a mouse renal ischemia–reperfusion model, Marschner et al¹⁴ used a ventilated heating chamber before and during surgery and maintained rectal core temperature within the 36.5-38.5 °C target for 99.3% ± 1.5% of the anesthetic period, compared with 0.7% ± 1.1% when only intraoperative heating was used. All these findings, together with the evidence linking peri-anesthetic hypothermia to experimental variability, highlight the need for simple, reliable warming solutions that act at induction and integrate smoothly into routine workflows.¹⁰

Despite the evidence supporting intervention during anesthetic induction, practical options remain limited. Commercial heated induction chambers are being introduced into the market, but they remain scarce and expensive. Alternative solutions, such as using heating blankets placed beneath the induction chambers, have been proven effective, but the application of this method can be inconsistent due to design variation (in both the heating element and the induction chamber) and may require extensive prewarming time to reach an effective temperature.¹

To address these limitations, we developed a cost-effective, Arduino-controlled heated induction chamber tailored for its use on mice. The system is designed to deliver precisely controlled, even, rapid heat and is built using widely accessible components. We validated its performance through bench-testing of safety and thermal accuracy and conducted a pilot study to assess its effectiveness in preserving normothermia during anesthesia induction.

Materials and Methods

Heated induction chamber design.

The heated induction chamber was designed to provide consistent and precise warming during the anesthesia induction phase in mice. The induction chamber measured 140 × 40 × 90 mm externally, yielding an estimated internal volume of ∼504 cm³, which accommodated a single adult mouse with sufficient space while minimizing excess dead space for anesthetic gas. At the core of the system was an Arduino Nano microcontroller (Arduino, Boston, MA), programmed to regulate temperature using a proportional integral derivative (PID) algorithm. This algorithm allowed the device to anticipate and correct temperature deviations smoothly, minimizing overshoot and undershoot while maintaining a stable setpoint.

Temperature was sensed by a 10 kΩ negative temperature coefficient (NTC) beaded thermistor embedded between a 100 × 150 × 2-mm aluminum plate and the 5-mm solid acrylonitrile-butadiene-styrene (ABS) chamber floor. The thermistor was positioned at the center of the chamber floor area. The aluminum plate served as both a thermal mass and a heat spreader, distributing warmth evenly across the chamber. Two 12-V, 20-W silicone heating pads were fixed beneath the plate using high-conductivity thermal paste to ensure efficient heat transfer.

The chamber’s walls were made from laser-cut acrylic, providing visibility and a smooth interior surface for easy cleaning. The housing and base, which secures the heating plate, electronics, and wiring, were 3D printed using ABS filament (Overture 3D Technologies, Houston, TX) on a Bambu Lab P1P printer (Bambu Lab, Austin, TX). This material was chosen for its thermal stability and mechanical strength. The base included an insulating base layer to minimize heat loss to the benchtop and a compartment for electronic components.

Heated induction chamber assembly.

The system was powered by a 12-V, 5-A DC power adapter connected to the heating elements through an IRLZ44N logic-level metal-oxide-semiconductor field-effect transistor (MOSFET), which acts as a high-efficiency switch controlled by the Arduino. The operator can set the desired temperature in the logic code, with the current setpoint and actual plate temperature displayed on an LCD screen. A red LED indicates active heating, and a momentary push-button provides reset or mode-change functionality (Figure 1).

Figure 1.

Prototype of the Arduino-Based Heated Anesthesia Induction Chamber. The system displays the programmed set temperature (40 °C) and the real-time base-plate temperature on an LCD screen during a bench-top performance test. A small breadboard hosts the power button and an LED indicator that illuminates while the heating elements are active.

All wiring with current <1 A was prototyped on a breadboard using jumper wires; other wiring with current >1 A was secured with female spade connectors outside the breadboard for reliable and safe electrical attachment of the heating pads and power terminals. A DC terminal adapter was mounted on the housing for quick connection to the power supply. The use of modular connectors and a compact microcontroller allows easy disassembly for maintenance or replacement of components.

Arduino control logic and PID regulation.

For readers less familiar with electronics, the Arduino functions as the ‘brain’ of the system, continuously measuring the temperature from the thermistor, comparing it to the target temperature to adjust the power delivered to the heating pads.

The PID control method acts as a ‘smart thermostat’ that not only responds to current temperature errors (proportional control) but also considers past trends (integral control) and anticipates future changes (derivative control). This prevents the system from cycling wildly between too hot and too cold and avoids the temperature ‘overshoot’ that can occur with simple on/off control.

In this application, the Arduino reads the thermistor temperature multiple times per second, calculates any difference from the setpoint, and modulates the MOSFET’s output using pulse-width modulation. The pulse-width modulation signal adjusts the effective voltage to the MOSFET gate, 0 V to turn off and ∼5 V to activate it, which allows the current to flow from the power source to the heating pads, fine-tuning the heat output to maintain stability. The LCD provides real-time temperature feedback to the user (Figure 1).

Safety assessment.

Surface temperature verification.

To ensure that the chamber floor achieved and maintained safe, effective warming conditions during both the prewarming and induction phases, we placed a secondary 10 kΩ NTC beaded thermistor directly on the chamber’s floor surface, where the animal has direct contact during induction. Both embedded and surface thermistors were connected to the Arduino to simultaneously log each temperature reading in the same manner to compare readings during the surface temperature validation.

Each bench trial began at ambient temperature, and heating was initiated until a stable temperature plateau was reached. The temperature difference (ΔT) between the embedded thermistor (used by the Arduino-based PID controller for regulation) and the surface thermistor was calculated to assess heating performance and duration of 3 working phases (illustrated in Figure 2):

• 1.Pre-warming phase, spanning from device startup until the maximum ΔT embedded surface (peak prewarming differential) was observed.
• 2.Stabilization phase, the first time interval in which both probes fluctuated within ±0.2 °C for at least 30 seconds.
• 3.Steady-state operation phase: the point after stabilization in which ΔT values remained consistent, with low variability (within ±0.1 °C) across repeated measurements for at least 2 minutes.

Figure 2.

Embedded and Surface Thermistor Temperatures (°C) over Time (min:s) During Bench Testing, Illustrating Prewarming, Stabilization, and Steady-State Phases.

This assessment aimed to quantify the relationship between the device’s internal temperature reading and the actual contact surface temperature that the animal would experience. We intended to evaluate whether the PID settings effectively prevent excessive overshoot (to eliminate any risk of surface overheating that could lead to animal harm) and provide a stable, constant surface temperature. The goal was to optimize the temperature setting to obtain a safe but efficient surface temperature of 37.5 ± 0.5 °C.

Infrared thermal imaging and heat distribution assessment.

To evaluate the uniformity of surface heating and detect potential hot or cold spots, we used a FLIR ONE Pro thermal imaging camera (Teledyne FLIR, Wilsonville, OR) to image the chamber base once the chamber reached steady-state operation. The infrared image provides a general heat distribution pattern, but resolution and accuracy limitations led to reduced precision. To address this, we manually overlaid a 5 × 6 grid (30 equal 20 × 25-mm sectors) on the ABS chamber floor, then measured the temperature of each sector using an infrared thermometer (model 96451; Cen-Tech, Camarillo, CA) to quantify the spatial temperature distribution. This analysis covered only the 100 × 150-mm central region, corresponding to the mouse’s direct contact area inside the chamber. The collected values were processed in Python 3.11 to generate a thermal map, in which the lowest and highest recorded temperatures defined the limits of a perceptually uniform color scale (magma colormap: dark purple/black to bright yellow), with intermediate values linearly interpolated across the grid to produce a smooth, continuous representation of the surface temperature field. Temperature deviations greater than ±0.5 °C from the mean were flagged as nonuniformities and considered for potential design adjustments.

Efficacy assessment.

Animal study design.

All animal procedures described in this study were reviewed and approved by the IACUC at the University of Arizona before the study execution. The University of Arizona’s animal care program is accredited by AAALAC International and adheres to all applicable federal and state regulations, as well as the Public Health Service Policy on Humane Care and Use of Laboratory Animals.

A paired, repeated-measures design was used in which each animal served as its own control, reducing the number of animals required. Ten animals (5 males, 5 females) were used in a within-subject design in which each animal underwent both experimental conditions, resulting in n = 10 paired observations. An a priori power analysis for a 2-sided paired t test (α = 0.05, target power = 0.80) to detect a 1.0 °C mean within-animal difference in core temperature, assuming an SD of paired differences of 0.8 °C, indicated that a total of 10 animals (5 per sex) was sufficient. An equal representation of males and females was selected and considered in our power analysis to avoid sex bias, but sex was not analyzed as a variable in this study. Ten adult C57BL/6 mice (males, 32.4 ± 3.2 g; females, 26.8 ± 4.8 g) from in-house breeding colonies were used. All animals were healthy and ∼20 to 22 weeks old at the time of study. All animals underwent 2 anesthetic conditions, in randomized order:

• 1.Nonheated induction chamber: the custom-built chamber was operated without the heating system activated (surface temperature = ambient).
• 2.Heated induction chamber: the custom-built chamber was preheated to an average surface temperature set at 37.5 °C.

The experimental unit was a single animal. A 7-day washout period was used between conditions. To minimize confounding, the order of animal testing within each condition was randomized. All animals first underwent the nonheated induction on the same day, and after the washout period, all underwent the heated induction on a single day, again in randomized order. This fixed order was intentionally selected for methodological reasons. Because the study used a within-subject design in which each mouse served as its own control, the risk of group-level confounding from nonrandom assignment was substantially reduced. Implementing alternating heated and nonheated inductions within the same day would have required repeatedly shifting the chamber from ambient temperature to ∼38 °C. Because the chamber retains heat for several minutes after power-off, this approach would introduce thermal carryover and risk contaminating ‘control’ exposures with residual warmth. In addition, frequent temperature transitions could have prolonged inter-animal intervals and altered the waiting-time environment, introducing additional handling- and stress-related variability.

Maintaining a fixed order, combined with a 7-day washout period, ensured consistent environmental conditions, chamber preparation, and workflow across all animals while minimizing uncontrolled day-to-day variability. Blinding was not used during experimental data acquisition, as the heated chamber had to be turned on and was designed with LED lighting indication when heating was active.

Inclusion and exclusion criteria.

All animals were included in the analysis, and no exclusion criteria were applied. No animals or data points were excluded.

Outcome measures.

The primary outcome measure was the core body temperature obtained from intraperitoneally implanted RFID temperature sensors, and secondary outcomes included physiologic parameters monitored during the maintenance phase (SpO₂, HR [heart rate], RR [respiratory rate], rectal temperature) and anesthesia recovery time.

Experimental animals and housing.

Mice were obtained from in-house breeding colonies. Animals were group-housed in individually ventilated cages (Lab Products, Seaford, MD) containing virgin paper pulp cellulose bedding (ALPHA-dri; Shepherd Specialty Papers, Watertown, TN). Enrichment items included a cardboard hut (Shepherd Shack; Shepherd Specialty Papers, Watertown, TN) and 2 types of nesting material: cotton square (Nestlet; Ancare, Bellmore, NY) and crinkled paper (Bed-r’Nest; The Andersons, Maumee, OH). Animals had free access to centralized reverse-osmosis water and NIH-31 modified open formula mouse diet (7913; Inotiv, West Lafayette, IN). The housing room was maintained at 20 to 26 °C and 30% to 70% relative humidity on a 12-hour light/12-hour dark cycle.

Experimental procedures.

RFID sensor implantation.

Seven days before the first anesthesia session subjected to the experimental variable, the mice were briefly anesthetized with isoflurane in oxygen for intraperitoneal implantation of an RFID temperature sensor (model UC-2112; Unified Information Devices, Brookfield, WI). Hair was clipped on the lower right abdominal quadrant, and the site was aseptically prepared with alternating povidone-iodine and isopropyl alcohol scrubs. The sensor, preloaded in a sharp 2.59-mm-diameter (∼13-gauge) cannula, was implanted through the prepared site using the manufacturer’s handheld insertion device. The insertion site was secured closed with a drop of tissue adhesive (Vetbond; 3M, St. Paul, MN). Mice were maintained on half-on/half-off heating pads and monitored until full recovery. Carprofen (10 mg/kg SC) was administered before anesthesia induction and every 24 hours for 2 days postoperatively.

Induction and maintenance of anesthesia.

Before each individual induction, the average chamber floor surface temperature was recorded for each condition. The average chamber floor temperature was obtained by averaging infrared thermometer (model 96451; Cen-Tech, Camarillo, CA) readings from the 4 corners and the center of the 100 × 150-mm camber inner surface. Mice were gently placed into the chamber, and anesthesia was induced with 4% isoflurane in oxygen at 1.5 L/min for 3 minutes. During induction, waste anesthetic gas was scavenged using a passive activated charcoal canister (Omnicon/F-AIR; A.M. Bickford, Wales Center, NY) attached to the chamber outlet. Before opening the chamber to remove the animal, the chamber was flushed with oxygen for ∼2 to 3 seconds to reduce residual isoflurane and minimize operator exposure. RFID core temperature readings were obtained every 30 seconds using a handheld reader (Unified Information Devices, Brookfield, WI).

Following induction, mice were transferred to a preheated multiparametric surgical platform (MouseMonitor S; Indus Instruments, Webster, TX) with the pad temperature set at 38 °C. Mice were positioned in ventral recumbency with all 4 paws gently secured with laboratory tape to the ECG pads of the monitoring platform to ensure accurate signal acquisition. Physiologic parameters, including SpO₂, HR, RR, and rectal temperature, were monitored and recorded every 30 seconds throughout the 10-minute maintenance phase. During maintenance, anesthesia was delivered as 2% isoflurane in oxygen at 700 mL/min, and RFID readings were obtained every 30 seconds.

At the end of the 10-minute maintenance period, isoflurane was discontinued, and oxygen flow was maintained until recovery. The tape securing the paws was gently removed; however, because the animals remained in ventral recumbency under the nose cone, return of the righting reflex could not be reliably assessed without handling. Therefore, recovery time was defined as the interval from cessation of isoflurane (with immediate flushing of the line with O₂) to the first purposeful movement, typically characterized by limb withdrawal from the pads or active pushing or shifting away from the nose cone. Once ambulatory, mice were returned to their home cages, where they remained half on, half off a heating pad for 60 minutes, and monitored until normal behavior resumed, and prior to returning to their housing IVC rack.

Statistical methods.

The experimental design included the induction chamber condition (heated compared with nonheated) as the independent variable and core body temperature as the primary dependent variable. Physiologic parameters (HR, RR, rectal temperature, and SpO₂) were also recorded to assess anesthetic stability but were not included in the statistical analyses, as the study was only powered to detect differences in core temperature between conditions Therefore, we carried out 2 separate repeated measures ANOVAs with induction method (nonheated compared with heated) as a within-subjects fixed factor and time (core body temperature at 30-second intervals) as another within-subjects fixed factor.

The first repeated measures ANOVA tested the change in core body temperature during the induction phase (0:00-3:00 minutes). The second repeated measures ANOVA tested the change in core body temperature during the maintenance phase (0:00-10:00 minutes). If significant main and interaction effects were detected, we followed up with preplanned comparisons between core body temperature at baseline (ie, 0:00 serves as control) and core body temperature at each subsequent 30-second interval. Preplanned comparisons were carried out for each induction method, which focused on detecting changes in core body temperature across time. We did not carry out comparisons between induction methods because our primary objective focused on determining the core body temperature that was reached at the end of the induction phase, and that value remained stable throughout the maintenance phase. Statistical significance for all analyses was set at α = 0.05 (SPSS, Chicago, IL). Exact P values are provided in the Results, and corresponding significance levels are denoted in figures using conventional symbols. Time-series data are expressed as mean ± SEM, with significant deviations from baseline highlighted at individual time points.

Results

Surface temperature validation and thermal stability.

The bench test condition that produced a surface temperature closest to the target of 38 ± 0.5 °C was achieved when the chamber’s internal set point (measured by the embedded thermistor) was set to 43 °C. Starting from an ambient temperature of ∼27 °C, the largest difference between the embedded and surface probes (ΔT) was reached ∼5 minutes (4:52 minutes) after the device was turned on, when the embedded temperature was 40.72 °C, and the surface temperature was 31.81 °C (ΔT = 8.91 °C). The system then gradually equilibrated as heat transferred from the internal heating elements to the chamber floor surface (Figure 2).

The first stabilization point (defined as both probes fluctuating by no more than ±0.2 °C during a 30-second interval) was reached ∼9 minutes (8:58 minutes) after starting the system. At this time, the embedded probe read 43.01 °C and the surface probe 37.38 °C, yielding a ΔT of 5.63 °C.

The chamber then entered steady-state operation, during which ΔT remained highly consistent over time, averaging 5.02 °C with very low variability (SD of 0.07 °C, range of 4.91 to 5.11 °C).

Infrared thermal imaging confirmed that surface heating of the chamber floor was generally uniform during the steady-state operation phase, although localized variations were present. Measurements obtained with a handheld infrared thermometer across a 5 × 6 grid traced on the ABS chamber floor ranged from 36.21 °C to 39.11 °C, with a mean of 37.53 °C (SD of 0.66 °C) when the embedded thermistor was set to a target of 43 °C. Eleven of the 30 measurement points (36.7%) deviated by more than ±0.5 °C from the mean, with 5 values above and 6 below, primarily located at the periphery of the chamber floor. Despite these deviations, the surface remained within a safe and physiologically appropriate range for mouse anesthesia induction, and no large hot or cold spots were detected.

For visualization purposes, Figure 3 presents the Python-generated thermal map derived from these discrete measurements. In this approach, the lowest and highest recorded values were assigned to the extremes of a perceptually uniform color gradient (magma colormap: dark purple/black to bright yellow), with intermediate values linearly interpolated across the grid to provide a smooth, intuitive heat map that conveys the subtle spatial differences in surface temperature.

Figure 3.

Python-Generated Thermal Map of the Chamber Floor Surface During Steady-State Operation with the Internal Temperature Set to 43 °C. Temperatures (°C) are shown within each sector. Values in blue are >0.5 °C below the mean (37.53 °C), and values in red are >0.5 °C above.

Animal study outcomes.

For the main outcomes of this section, that is, core body and rectal temperatures, the results are expressed as mean ± SEM to better reflect the precision of the mean at each time point and to facilitate clear graphical presentation of repeated measures across time. All other data (surface temperature, physiologic data, and anesthesia recovery time) are reported as mean ± SD.

During the efficacy assessment, the chamber surface temperature during the unheated (control) condition averaged 24.23 ± 0.25 °C, closely matching ambient room temperature and exhibiting minimal variability across iterations. During the heated induction condition, the average surface temperature was 37.65 ± 0.54 °C, with slightly greater variability but showing consistent performance across iterations and alignment with the intended warming target of 37.5 °C.

During postimplantation monitoring of intraperitoneal RFID temperature transponders, abnormally low readings were occasionally observed in male mice. Closer examination revealed that the transponders, although implanted in the abdominal cavity, had migrated into the scrotum in some animals. Their presence in the scrotum was readily detected by palpation and confirmed radiographically (Figure 4). Gentle manual manipulation was sufficient to reposition the transponder into the abdominal cavity, after which temperature values immediately returned to physiologically normal ranges. On experimental days, when abnormally low baseline (preinduction) temperatures were detected, transponders were gently repositioned before beginning the induction phase to ensure that time 0 and subsequent measurements accurately reflected true intraperitoneal core body temperature.

Figure 4.

Radiographs of Mice Implanted with Intraperitoneal RFID Temperature Microchips. In (A) (dorsoventral), a transponder in the abdominal cavity of one animal is shown, whereas (B) (dorsoventral) and (C) (laterolateral) show a transponder that migrated into the scrotum of a different animal.

At baseline, core temperatures were nearly identical across conditions (control, 36.97 ± 0.24 °C; heated induction, 37.01 ± 0.14 °C; P = 0.888). During the induction phase, significant main effects of time (P < 0.001), induction condition (P = 0.001), and their interaction (time × condition, P < 0.001) were observed. When animals underwent the unheated (control) condition, body temperature declined rapidly, reaching 35.38 ± 0.27 °C at 3:00 minutes. In contrast, during the heated induction condition, temperature increased steadily, reaching 37.82 ± 0.26 °C at 3:00 minutes (P < 0.001). Within-animal comparisons relative to baseline (0:00 minutes) indicated that temperature decreases in the control condition became significant by 1:00 minutes (P = 0.004) and remained significant at all subsequent time points (all P < 0.001; Figure 5). In the heated induction condition, temperature increases emerged at 0:30 minutes (P = 0.020) and persisted through the end of the induction phase (all P < 0.05; Figure 5).

Figure 5.

Mean ± SEM Core Body Temperature (n = 10) During the 3-min Induction and 10-min Maintenance Phases. Significant main effects of time, condition, and their interaction were detected (P < 0.001). Symbols indicate differences from baseline: *P ≤ 0.05, ^†P ≤ 0.01, and ^‡P ≤ 0.001.

During the 10-minute anesthetic maintenance phase, significant main effects of time (P < 0.001), induction condition (P < 0.001), and their interaction (P < 0.001) were also detected. Throughout this period, mice maintained higher core temperatures under the heated induction condition compared with the unheated condition. When unheated, animals gradually rewarmed from 35.68 °C at 0:00 minutes to 37.43 °C at 10:00 minutes but remained consistently below the temperatures observed under heated induction, where normothermia was maintained throughout. Simple contrast analyses showed that temperature stabilized by 6:00 minutes in the control condition (P = 0.111) and remained stable across all time points in the heated condition (all P > 0.05), with values of 37.80 ± 0.16 °C at the start and 37.85 ± 0.16 °C at the end of the maintenance phase.

Core body temperature compared with rectal temperature.

Concurrent measurements obtained during the anesthetic maintenance phase demonstrated the expected systematic differences between intraperitoneal RFID-derived core temperatures and rectal probe temperature values. In both experimental conditions, rectal measurement readings were consistently lower than core intraperitoneal temperatures. During the unheated (control) condition, rectal readings were on average 1.18 ± 0.53 °C lower than core values at each matched time point, while during the heated induction condition, rectal temperatures were also lower than core values, but with an average difference of 0.87 ± 0.41 °C (Figure 6).

Figure 6.

Mean ±SEM Intraperitoneal (Core) and Rectal Temperatures of Mice During Anesthetic Maintenance Under Control (Unheated) and Heated Induction Conditions. Core temperature was recorded via intraperitoneally implanted RFID transponders, and rectal temperature was simultaneously measured with a rectal probe.

Physiologic stability during anesthesia.

HR values during the maintenance phase showed some within-animal variability across conditions. During the unheated (control) condition, HR averaged 435 ± 83 bpm, while during the heated induction condition it averaged 475 ± 108 bpm. Despite the trend toward higher mean values in the heated induction condition, no systematic cardiac instability was evident under either condition.

RR during the maintenance phase also showed some within-animal variability across conditions. During the unheated (control) condition, RR averaged 119 ± 32 breaths/min, while during the heated induction condition, RR averaged 126 ± 32 breaths/min. Despite this modest difference in central tendency, no meaningful differences were observed. Although RR deviations >25% of the mean occurred under both conditions, RR values remained within physiologically acceptable ranges throughout.

During the initial minutes of anesthetic maintenance, not all animals contributed physiologic data to every time point because proper instrumentation (paw taping to ECG electrode, ECG transducer gel application, and rectal probe positioning) required from several seconds in some animals to up to a few minutes in others to achieve a stable signal and reliable measurements. These procedures occurred concurrently with continuous intraperitoneal temperature acquisition (the primary outcome), so brief gaps in the HR and RR recordings resulted. Figure 7 indicates the number of animals represented at each depicted time point. Only one side of the SEM is shown to avoid overlap between conditions and to facilitate visualization of temporal trends. Importantly, despite these early gaps, the data demonstrate no evidence of systematic cardiorespiratory instability under either condition.

Figure 7.

Mean ± SEM Temporal Profiles of Heart Rate (A) and Respiratory Rate (B) During Anesthetic Maintenance Under Control (Unheated) and Heated Induction Conditions (n = 10). Only one SEM bar is shown per time point to minimize overlap. Numbers above selected points indicate time points with <10 animals due to instrumentation delays during early data collection, or signal loss.

SpO₂ remained within physiologically acceptable ranges during both conditions throughout the anesthetic maintenance phase. During the unheated (control) condition, SpO₂ averaged 93.4% ± 7.6%, while during the heated induction condition, SpO₂ averaged 92.9% ± 4.9%. Although both conditions maintained average values above the hypoxemic threshold (90%), SpO₂ during the unheated condition trended slightly higher. All animals remained near these values, with only occasional transient dips observed. No consistent desaturation below the hypoxemic threshold was observed under either condition.

The average anesthesia recovery time (time from isoflurane discontinuation and system flush to first voluntary movement) was also nearly identical across conditions: 1:36 ± 0:29 minutes (96.4 ± 29.25 seconds) following the unheated (control) condition and 1:28 ± 0:21 minutes (88.3 ± 21.41 seconds) following the heated induction condition (Table 1).

Table 1.

Physiologic Parameters During Anesthetic Maintenance and Recovery Time in Mice Exposed to Both Experimental Conditions: Control (Unheated) and Heated Induction in a Repeated-Measures Design

Parameter	Control (mean ± SD)	Heated induction (mean ± SD)
Heart rate, beats/min	435 ± 83	475 ± 108
Respiratory rate, breaths/min	119 ± 32	126 ± 32
SpO2, % hemoglobin saturation	93.4 ± 7.6	92.9 ± 4.9
Time to first movement, min:s	1:36 ± 0:29	1:28 ± 0:21

Discussion

Induction chamber function.

In terms of function and reliability, our designed chamber reached a stable operating temperature within 10 minutes and provided a homogeneous heated surface. The first stabilization point was reached at 8:58 minutes after system activation. At that time, the device reached a steady state and maintained consistent performance throughout all experimental sessions. These observations are promising when compared with a recent study that showed a commercial heated induction device and a cost-effective alternative (a heating pad placed beneath a standard induction chamber) were both preheated for 2 hours to achieve approximately 28 °C.¹ It is unclear whether the commercial device required this extended preheating period, or whether it was held for that duration to match the slower performance of the alternative system, but regardless, our device required less than 10 minutes to stabilize at a considerably higher target temperature (approximately 38 °C). We consider this warm-up and stabilization time to be compatible with all standard mouse anesthetic practices and readily adoptable in most laboratory settings.

Once stable performance was established, we next considered the physiologic relevance of the target surface temperature. We decided to set the chamber surface temperature to 37.5 °C, a target temperature within the generally accepted normal body temperature range in mice (36.5-38 °C).^1,15 This target is consistent with contemporary rodent anesthesia clinical practice,⁴ and it also aligns with values reported across anesthetic studies, where investigators typically aim to maintain mice within a body temperature of 36 to 38.5 °C to prevent hypothermia and ensure stable physiology.^3,5,12,16 Maintaining body temperature within this range during anesthesia is considered essential, as even modest deviations can alter anesthetic depth, cardiovascular and respiratory stability, and quality of recovery.^2,8 Recent large-scale anesthetic physiologic impact evaluations across substrains contribute to emphasizing the need for careful physiologic monitoring,¹⁷ where thermal support and stability during anesthesia represent a key element of success.

For our in vivo study, the prototype was configured with only the embedded thermistor to avoid introducing obstacles or elements that might injure the animals during induction. To achieve the desired average surface temperature of approximately 37.5 °C (0.5 °C below the validated surface temperature to minimize inadvertent thermal injuries), the thermistor set point was maintained at approximately 42.5 °C. Under these conditions, the device performed reliably, and sequential use for multiple animals without powering down did not compromise thermal stability performance. Additional verification with a handheld infrared thermometer prior to each animal use confirmed that the chamber floor surface remained within the target operating range, with an average of 37.65 ± 0.54 °C across 10 consecutive trials. In total, the device was connected and performed at a steady working state for approximately 4 hours 30 minutes. During the validation bench testing runs, the target internal temperature was set at 43 °C, and the average working temperature measured by the surface thermistor was approximately 38 °C, consistently 5.0 °C cooler than the embedded thermistor reading. This offset, however, should not be generalized, as it reflects the specific characteristics of the current design. The observed offset is heavily determined by key design-specific factors such as the thermal conductivity of the primary material used (ABS, 0.17-0.25 W/m·K), the chamber floor thickness (5 mm), and the validated internal set temperature (43 °C). Alterations to any of these parameters would be expected to produce a different temperature differential, an aspect we plan to investigate further as we adapt the prototype for use in rodent species of larger size (eg, rats).

Heated anesthesia induction: safety and efficacy.

This animal study demonstrates the safety and efficacy of our custom-built, heated induction chamber specifically designed for mice. During the heated induction condition, mice maintained normothermia throughout the induction period and even exhibited a modest rise in core temperature, whereas during the unheated control condition, the same animals experienced a rapid and clinically relevant decline.

Our results confirm that a heated induction chamber is an effective strategy to prevent the rapid hypothermia typically observed during the early stages of anesthetic induction with isoflurane in mice. These findings align with prior reports in both mice^1,3 and rats,^6,18 demonstrating that induction warming and prewarming strategies mitigate the steep temperature decline that occurs in rodents during inhaled anesthetic induction. Some of the strategies already described in rodents include prewarming through heated forced air, either aimed at increasing core body temperature to a specific target^6,18 to provide a controlled heated environment,^3,9 or a heated induction chamber in which the contact surfaces are warmed.¹ The mechanisms underlying the protective effect of active warming are well established, counteracting the redistribution of warm core blood to peripheral tissues that normally accounts for about 80% of the initial hypothermia during anesthesia. Although mice can exhibit short-term fluctuations of approximately 1 °C within their narrow (1-3 °C) thermoneutral zone,² under anesthesia (when the behavioral and physiologic compensation mechanisms are abolished or impaired), this can represent a physiologically meaningful deviation. If maintained during the anesthetic procedure, perioperative hypothermia is associated with adverse outcomes, such as delayed recovery and increased complication risks.² Providing a warm environment and a warm contact surface immediately upon loss of consciousness can significantly minimize this initial temperature loss.

Although our study evaluated this approach only in the context of inhaled anesthesia, the findings suggest that a heated induction chamber could also be of value when injectable protocols are used in mice. Significant hypothermia is a well-documented complication of injectable regimens, including ketamine–xylazine and related combinations, which can rapidly depress cardiovascular and thermoregulatory function if not supported.^5,13,16 Previous reports demonstrate that warmed induction boxes effectively mitigated hypothermia in rats undergoing injectable anesthesia,⁶ supporting the premise that our heated chamber could provide comparable benefits in mice by preventing temperature loss during the vulnerable induction period. Recent work in the field has shown that mice physiologic responses under injectable anesthesia can be influenced not only by drug selection but also by substrain and sex.¹⁷ In this context, standardized supportive care during anesthesia induction can help reduce confounding factors. Placing the animals in a warmed environment immediately after injection (before the loss of the righting reflex) could help mitigate the early redistribution hypothermia that occurs during anesthetic induction, improving physiologic stability and recovery quality.

In addition to its performance in terms of safety and efficacy, the developed device is inexpensive to build (material cost < $250), enhancing its feasibility for broad adoption as a practical refinement in rodent anesthesia.

Comparative heating device performance.

A variety of approaches, including circulating water blankets, heating pads, gel packs, and reflective foils, have been described and tested for their ability to mitigate hypothermia during rodent anesthesia; however, their effectiveness varies considerably across studies and devices.^3,5,11,19 In our study, control animals returned to baseline normothermia relatively quickly (>3 minutes) once placed on a preheated surgical platform. In contrast, in the work of Gaskill and colleagues,¹ control mice regained only part of their lost body temperature and remained below the normal physiologic range throughout the 10-minute maintenance phase, while even the heated groups demonstrated modest declines during comparable anesthetic intervals. These differences likely reflect the lower set temperature of the water blanket used in that study (33 °C during the maintenance phase) and the inherent variability of circulating warm water systems, which have been shown to require prolonged warm-up times and often fail to reach or maintain their user-set values.¹¹ Blanket design features, such as the pattern configuration and diameter of the water channels, may also limit the effective contact area with the small body surface of mice, further reducing the efficiency of heat transfer.

In contrast, surgical platforms such as the one employed in our study deliver consistent surface temperatures through a more uniform body contact, which together can help explain the faster and more complete recovery of core body temperature observed in our animals. Although conventional warming devices used during the maintenance phase can eventually stabilize body temperature, preventing the sharp decline at induction, as achieved with our chamber, may offer more robust protection against the physiologic consequences of peri-anesthetic hypothermia and could reduce the need for prolonged postoperative heating supplementation, contributing to minimizing postanesthetic hypothermic events.¹³

Temperature monitoring considerations.

Temperature monitoring using implantable devices in rodent studies has been performed using both subcutaneous^1,3,5 and intraperitoneal^6,9,13 locations. Subcutaneous implantation is minimally invasive and widely adopted; it is inherently more superficial and, therefore, in our opinion, more easily influenced by positional effects and proximity to external heat sources. This can lead to overestimation or underestimation (depending on the animal’s positioning) of true core temperature. In contrast, intraperitoneal placement offers a direct and physiologically accurate assessment of core body temperature, better reflecting the thermoregulatory status under anesthesia. For this reason, our study employed intraperitoneal RFID transponders aiming for robust measurements of core temperature dynamics during anesthetic induction and maintenance.

Concurrent measurements obtained during the anesthetic maintenance phase demonstrated a strong correlation⁵ and systematic differences between intraperitoneal RFID-derived core values and rectal probe readings. In both experimental conditions, rectal temperatures were consistently lower than intraperitoneal core measurements. In the control condition, rectal readings averaged 1.18 ± 0.53 °C below core values at each matched time point, while during the heated induction condition, the gap was slightly smaller but still evident, averaging 0.87 ± 0.41 °C (Figure 6). We think that this observation is highly relevant given that rectal temperature remains the most commonly used method to approximate and control core temperature in rodent anesthetic studies^16,17 and the standard in laboratory animal practice.⁴

Because many heating systems rely on rectal probe feedback to cycle warming devices on and off, the ∼1 °C gap between rectal and core values must be carefully considered. If a device is programmed to maintain a rectal temperature target of 37.5 °C, the true intraperitoneal temperature according to our data may reach 38.5 °C or higher, placing the animal at high risk of inadvertent hyperthermia. Maintaining physiologic normothermia in rodents requires precise monitoring, as even small deviations can prolong recovery or alter anesthetic depth.^6,9,13 Thus, we find that relying solely on rectal temperature without accounting for this offset could inadvertently push animals beyond the upper physiologic range.

A final important observation related to temperature monitoring in this study was the migration of intraperitoneally implanted RFID transponders into the scrotal sac of male mice. This circumstance, anatomically feasible due to the permanently open inguinal canal in this species, has not, to our knowledge, been previously reported. Although the displacement was readily corrected by gentle digital manipulation (after which temperature values immediately returned to physiologic ranges), its occurrence can have important implications for data quality if undetected. Unrecognized migration could lead to significantly low or erratic readings, potentially confounding experimental outcomes. This finding emphasizes the need for researchers to confirm transponder location in male mice prior to data collection. Noninvasive strategies include checking baseline values, gentle scrotal palpation, or radiographic confirmation when available. Being aware of this phenomenon is essential for ensuring accurate temperature monitoring and maintaining reproducibility in studies that rely on small intraperitoneal RFID transponders in rodents.

Maintenance phase physiology.

During the anesthetic maintenance period, physiologic stability in our study was broadly comparable to recent reports of mice under inhaled isoflurane anesthesia with active thermal support.^16,17 SpO₂ remained consistently within clinically adequate ranges, and both HR and RR exhibited the expected interanimal variability while remaining within physiologic limits. Interestingly, HR and RR trended slightly higher in the heated induction group. These trends (and the expected early variability related to instrumentation) are visually evident in the temporal HR and RR profiles (Figure 7), which demonstrate stable cardiorespiratory function under both conditions, with the heated induction group remaining slightly but consistently above the control group for both parameters throughout most of the maintenance phase. Although these differences were subtle and not statistically analyzed, they may reflect increased metabolic activity or an early compensatory response to incipient hyperthermia, consistent with prior observations that body temperature modulates cardiovascular stability and respiratory drive in isoflurane-anesthetized mice.^4,5,20

Importantly, our findings align with those of recent studies demonstrating that early application of thermal support mitigates the rapid decline in body temperature otherwise observed in unheated induction chambers.¹ By maintaining higher core temperatures during the heated induction condition, mice showed physiologic stability without evidence of respiratory or hemodynamic compromise, reinforcing prior reports that effective warming reduces variability and improves recovery trajectories.¹³ Although our anesthetic exposures were shorter and our cohort was smaller than in some previously published work, the trends we observed are consistent: prewarming or heated induction blunts early heat loss, and sustained and effective postinduction warming during maintenance accelerates recovery from the hypothermia typical of standard induction protocols.

Limitations.

Our study has several important limitations to acknowledge. Although animals were continuously monitored during induction, we did not record the time to loss of the righting reflex, which would have provided an additional quantitative measure of anesthetic induction depth and progression. The study was conceptualized as a pilot and thus was powered to detect differences in core temperature, not secondary physiologic endpoints. As a result, the trends observed in HR and RR cannot be interpreted robustly and, therefore, we intentionally did not perform formal statistical analyses or sex-stratified comparisons for these variables. Nonetheless, the variability patterns observed in this pilot study for these physiologic measures provide useful preliminary estimates to inform sample-size calculations and guide the design of adequately powered follow-up studies for those secondary physiologic endpoints.

Another factor to consider is that we only evaluated a single strain (C57BL/6) of mice with one inhalant regimen under controlled conditions. To enable generalization of these findings to other strains, ages (juvenile/aged), body masses, and injectable protocols, adequately designed studies are needed. Also, while both sexes were included, the study was not stratified or powered to test sex-specific effects. Thus, larger, stratified cohorts are needed to determine whether males and females benefit differentially from heated induction and whether sex modifies the temperature–cardiorespiratory coupling observed.

It is also noteworthy that our study focused only on the induction and a short maintenance interval to assess the validity and efficacy of the custom chamber design. Extending monitoring through the postanesthetic period, such as the ones done by Gaskill et al,¹ would allow us to test whether the early thermal protection provided by our device translates into longer-lasting functional benefits.

Conclusions

This study confirms that heated anesthesia induction is a simple yet highly effective refinement for preventing early hypothermia in mice, with clear potential to improve both animal welfare and the reproducibility of experimental outcomes. The cost-effective heated induction chamber we describe in this study is built from accessible components and validated for safety and efficacy. Our heated induction chamber offers a practical, scalable tool that can be readily integrated into routine workflows. While our future work will assess its performance under diverse anesthetic regimens and procedural contexts, our findings position heated induction as a strong candidate for incorporation into new standards of refined rodent anesthetic practice.

Conflict of Interest

The authors have no conflicts of interest to declare.

Funding

This work was internally funded by the University Animal Care department at the University of Arizona. A pre-veterinary medicine student participating in the University of Arizona College of Veterinary Medicine Summer Scholars Research Program contributed to the development and validation of the Arduino-based heated induction chamber as part of the program, which provides stipend-supported summer research experiences for veterinary and pre-veterinary students.

Data availability

All data supporting the findings of this study are available from the corresponding author upon reasonable request.