Anesthesia Monitoring Parameters for Laboratory Animals

By Dr. Zubair Khalid, DVM, MS, PhD · 2026-08-11

Key Takeaways

• Species-specific physiological parameters and monitoring ranges are critical: Small laboratory mammals, particularly mice and rats, exhibit high metabolic rates and rapid heat loss, necessitating tailored monitoring. For instance, target rectal temperatures differ significantly between mice/rats (36.5-38.0 °C) and rabbits (38.0-39.5 °C), and hypothermia is a primary risk factor leading to prolonged recovery and bradycardia.
• Anesthetic depth assessment requires a multi-modal approach: Relying solely on reflexes like pedal withdrawal or palpebral response is insufficient due to species and strain variations; a composite assessment integrating reflex responses, respiratory pattern, heart rate trends, and mucous membrane color is essential. Loss of the righting reflex indicates unconsciousness but not surgical depth in rodents.
• Technical limitations of standard monitoring equipment in small patients are significant: Pulse oximetry probes designed for humans often fail to transmit light effectively through rodent tissue or produce motion artifact, leading to unreliable SpO₂ readings. Capnography in mice requires low dead space adapters and high sampling rates to avoid rebreathing and signal damping due to small tidal volumes.
• Hypothermia is the most consequential complication and requires proactive management: Continuous rectal or esophageal temperature measurement is crucial, and intervention is needed when temperature falls below species-specific baselines, not just predetermined absolute thresholds. Warming devices must be used cautiously to avoid thermal injury.
• Respiratory depression and bradycardia are common adverse events requiring early detection: Isoflurane can cause marked respiratory depression, while certain injectable agents like ketamine-xylazine combinations induce significant bradycardia. Early detection via capnography or direct observation, and continuous pulse oximetry or ECG respectively, is vital.
• Accurate documentation of anesthetic parameters is paramount for reproducibility and welfare: Anesthetic records must detail baseline values, agent administration, monitoring parameters at regular intervals (minimum every five minutes for stable patients), and any interventions. This supports clinical care, scientific validity, and regulatory compliance.

This article provides a practical framework for monitoring anesthesia depth and physiological status in common laboratory animal species, with emphasis on mice, rats, and rabbits. It is written for veterinary researchers and laboratory animal veterinarians who design, perform, or review anesthetic protocols in biomedical research. The content addresses the selection of monitoring parameters, the interpretation of derived values in small patients, and the species-specific limitations of standard monitoring equipment.

The central clinical question is how to distinguish adequate anesthetic depth from dangerously deep or inadequate planes when working with animals whose small size, high metabolic rate, and rapid heat loss distort familiar monitoring signals. A secondary question concerns the reporting standards expected in peer-reviewed publications, since incomplete anesthetic monitoring data compromises the reproducibility of experimental results. The National Research Council Guide for the Care and Use of Laboratory Animals establishes the institutional expectation that perioperative monitoring be appropriate to the species and procedure, and the NC3Rs resources on refinement provide practical guidance for reducing anesthetic morbidity through better monitoring.

At a Glance

Physiological Basis of Monitoring in Small Laboratory Mammals

The physiologic challenges of anesthetic monitoring in laboratory rodents stem from their high surface area to volume ratio, high basal metabolic rate, and small absolute blood volume. A mouse loses heat rapidly under anesthesia because thermoregulation is depressed and the body surface is large relative to mass. Hypothermia in turn reduces anesthetic requirement, slows drug metabolism, and depresses respiratory drive, creating a self-reinforcing cycle of deepening anesthesia. The review of anesthetic considerations in preclinical mouse research identifies hypothermia and hypoglycemia as the two most consequential complications in murine anesthesia, and both are detectable through routine monitoring if the observer knows what to measure.

Cardiorespiratory physiology in rodents differs from that of larger mammals in ways that affect monitor interpretation. Mice have a resting heart rate of 500 to 700 beats per minute and a respiratory rate of 100 to 200 breaths per minute, values that approach the response limits of some clinical monitors. The small tidal volume of a mouse, roughly 0.15 to 0.2 mL, means that capnography sampling lines can aspirate a significant fraction of the breath if flow rates are not adjusted. Pulse oximetry probes designed for human digits often fail to transmit light through rodent tissue or produce motion artifact that obscures the signal. These technical constraints, not the physiology itself, are the most common cause of unreliable monitoring data in mice.

Anesthetic Depth as a Continuum

Anesthetic depth is not a single state but a graded depression of reflex responses, motor tone, and autonomic function. The classic Guedel stages, developed for ether anesthesia in humans, do not translate directly to rodents because the reflex sequences differ and modern agents produce different patterns of depression. In practice, depth is assessed by testing responses that are lost in a predictable order. The pedal withdrawal reflex, palpebral reflex, and ear pinch response disappear at surgical planes in most laboratory species, while the corneal reflex and spontaneous respiratory effort persist until deeper planes. Loss of the righting reflex marks a light plane in rodents, but it is not sufficient for surgery.

The comparative study of vital signs during injectable and inhalant anesthesia in mice demonstrates that different protocols produce different dissociations between depth and vital signs. In that study, ketamine and xylazine combinations produced surgical anesthesia with marked bradycardia but relatively preserved respiratory rate and oxygen saturation. Medetomidine, midazolam, and butorphanol combinations produced lighter anesthesia with lower heart rate and SpO₂, but these changes reversed rapidly after atipamezole administration. Pentobarbital produced inconsistent depth, with 62.5% of mice failing to reach a surgical plane, and was associated with lower SpO₂. Isoflurane produced the most stable oxygen saturation but the greatest respiratory depression. These findings indicate that no single vital sign predicts depth across protocols, and that the monitoring plan must be tailored to the pharmacologic profile of the chosen agents.

Temperature Monitoring and Thermal Support

Body temperature is the single most important monitored parameter in small laboratory mammals because hypothermia affects every other physiologic variable. A 2 °C drop in core temperature reduces heart rate by roughly 10% to 20% in mice, depresses respiratory drive, prolongs recovery from injectable agents, and increases perioperative mortality. The anesthetic considerations review for preclinical imaging emphasizes that temperature control is particularly critical during imaging procedures, where animals are often positioned on cold scanner beds and cannot be easily wrapped or covered.

Rectal temperature probes are the standard method in rodents, but they must be inserted gently to avoid perforation and must be secured to prevent expulsion during recovery. Infrared thermometry of the ear or paw correlates poorly with core temperature in small species and should not replace rectal measurement. For rabbits, the rectal temperature is normally higher than in rodents, and the same probe systems work with appropriate probe size. Warming devices include circulating water blankets, forced air warmers, and heat lamps, but each carries a burn risk in small patients. A heat lamp positioned too close to a mouse can cause thermal injury before the rectal temperature rises detectably, so indirect heating with a circulating water blanket under the animal is generally safer.

Respiratory Monitoring

Respiratory rate is the most accessible respiratory parameter in small laboratory animals and can be measured by direct observation of thoracic wall movement, by impedance-based sensors, or by capnography. Direct observation is reliable in mice and rats because the chest wall movements are visible, but it requires the observer to remain attentive for a full minute to obtain an accurate rate. The systematic review of anesthesia reporting in high impact journals found that respiratory monitoring was inconsistently reported across experimental studies, which limits the interpretability of physiologic outcomes in published research.

Capnography in rodents requires low dead space adapters and high sampling rates to avoid rebreathing and signal damping. Sidestream capnographs with sampling rates of 50 to 150 mL/min can aspirate more gas than a mouse breathes in a minute, so mainstream or microstream analyzers are preferred. End tidal carbon dioxide values in rodents are typically 35 to 45 mm Hg at surgical planes, but the waveform shape is more informative than the absolute number. A plateau phase indicates adequate sampling, while a sloping or absent plateau suggests the sample is diluted by fresh gas flow or the tidal volume is too small for the analyzer.

Pulse oximetry is widely used but has important limitations in rodents. The vital signs comparison study in mice reported that SpO₂ values were stable under isoflurane but lower under medetomidine combinations, and that the readings were sensitive to probe placement and motion. In mice, the probe is typically placed on the hind paw or tail, and the signal is often lost during movement or vasoconstriction. A reading above 90% is generally reassuring, but a low reading should be confirmed by direct assessment of mucous membrane color and respiratory effort before adjusting the anesthetic plane.

Cardiovascular Monitoring

The cardiovascular system is the second pillar of intra-anesthetic assessment in laboratory mammals. Heart rate and pulse quality reflect anesthetic depth, nociceptive stimulation, and the direct cardiodepressant effects of anesthetic agents. In mice, injectable combinations produce markedly different cardiovascular profiles. Ketamine-xylazine causes remarkable bradycardia, whereas medetomidine-midazolam-butorphanol produces a relatively lower heart rate that reverses rapidly with atipamezole administration. These differences matter clinically because a heart rate that is acceptable under one protocol may indicate excessive depth under another.

Heart Rate Assessment

Auscultation with a pediatric stethoscope is feasible in rats, guinea pigs, and rabbits but unreliable in mice. Palpation of the precordial impulse or peripheral pulse is practical in larger laboratory mammals but gives only rate, not rhythm. Electrocardiography provides continuous rate and rhythm data and is the standard for detecting arrhythmias, which occur with hypoxemia, hypercapnia, and certain anesthetic agents. Pulse oximetry plethysmography offers a continuous heart rate trace and is widely used in mice, though motion artifact and poor peripheral perfusion degrade the signal.

Doppler ultrasound flow probes placed over a peripheral artery, most commonly the tail in rats or the medial saphenous in mice, provide an audible pulse and are more tolerant of low perfusion states than pulse oximetry. The Doppler signal confirms perfusion, also electrical activity, and is therefore more informative than electrocardiography alone when cardiac output is compromised.

Pulse Quality and Perfusion

Pulse quality is assessed by palpation of the femoral artery in rats and rabbits, the tail artery in mice, and the auricular artery in rabbits. A weak, thready pulse with tachycardia suggests hypovolemia or deep anesthetic plane. A bounding pulse with bradycardia may indicate excessive depth or a vagal response. Capillary refill time, assessed on the plantar surface of the foot or the gingiva, is prolonged beyond two seconds with poor perfusion, but this test is difficult to interpret in small patients with rapid normal circulation.

Blood Pressure Measurement

Direct arterial blood pressure measurement is the reference method but requires arterial catheterization, which is technically demanding in mice and rats. The carotid artery is commonly catheterized in rats for this purpose, while the femoral artery is used in rabbits. Direct measurement is indicated for procedures with significant blood loss, prolonged surgery, or when vasoactive drugs are used.

Noninvasive oscillometric and tail-cuff methods are available for rats and rabbits but are less accurate during hypotension and are affected by vasoconstriction from hypothermia and anesthetic agents. The tail-cuff method requires warming the animal and produces intermittent readings that may not capture acute changes. In mice, noninvasive blood pressure measurement is unreliable and is rarely used during anesthesia.

Oxygenation and Ventilation Monitoring

Pulse oximetry measures hemoglobin oxygen saturation and is the most widely used oxygenation monitor in laboratory animal anesthesia. In mice, the probe is typically placed on the hindlimb, tail, or neck, and the signal is affected by peripheral vasoconstriction, hypothermia, and probe positioning. The isoflurane group in one comparative study showed the most stable oxygen saturation among the protocols evaluated, despite a marked decrease in respiratory rate, indicating that stable saturation can coexist with significant respiratory depression.

Capnography measures end-tidal carbon dioxide and provides a continuous estimate of ventilation. Sidestream capnography with a low dead-space adapter is used in rats and rabbits, but the sampling flow rate can entrain room air and dilute the reading in very small patients. Mainstream capnography is impractical in mice. In rabbits, nasal cannula capnography provides a trend but underestimates arterial carbon dioxide. Capnography also confirms correct endotracheal tube placement and detects circuit disconnection, which is especially valuable during long imaging procedures where direct observation is limited.

Arterial blood gas analysis is the reference standard for ventilation and acid-base assessment but is invasive and provides only intermittent data. It is indicated when capnography is unavailable or unreliable, when there is suspected ventilation-perfusion mismatch, or when metabolic derangements are expected. In rabbits, arterial sampling from the auricular artery is straightforward. In rats, the femoral or tail artery is used, and in mice, arterial sampling is technically challenging and is usually reserved for terminal procedures.

Reflex Assessment and Anesthetic Depth

Reflex testing remains the primary method for assessing anesthetic depth in small laboratory mammals, particularly when electronic monitoring is limited. The pedal withdrawal reflex, elicited by firm toe pinch, is the most commonly used indicator. Loss of this reflex indicates surgical anesthetic depth in most species. The palpebral reflex, elicited by gentle touch of the medial canthus, is lost at a lighter plane and is useful for confirming that the animal is not too deep. The corneal reflex, elicited by gentle corneal touch, is lost only at deep planes and its absence signals impending anesthetic overdose.

The righting reflex is used during induction and recovery in rodents. Loss of the righting reflex indicates loss of consciousness but not surgical depth. In rabbits, the ear pinch reflex and pedal reflex are used, and the jaw tone is assessed by gentle opening of the mouth. The tail flick reflex in mice and rats is a spinal reflex that persists at surgical depth and is not a reliable indicator of consciousness.

Reflex assessment has limitations. It is subjective, requires repeated stimulation that can cause tissue trauma, and does not detect hypoxemia or hypercapnia before they become severe. Reflexes should be interpreted alongside physiologic parameters, and the absence of a pedal reflex should not be the sole criterion for proceeding with surgery if other parameters indicate instability.

Species-Specific Normal Ranges

Normal physiologic parameters vary substantially across laboratory species, and reference ranges must be interpreted in the context of the anesthetic protocol, the animal's age and strain, and the procedure being performed. The table below provides reference ranges for common laboratory species. These values are derived from standard veterinary references and should be used as guidelines instead of absolute thresholds.

Heart rates at the upper end of these ranges are common in conscious animals and during light anesthesia. A mouse heart rate below 300 beats per minute under isoflurane anesthesia may indicate excessive depth, whereas the same rate under ketamine-xylazine may be expected. Respiratory rates are similarly protocol-dependent. The marked decrease in respiratory rate seen with isoflurane in mice requires vigilance, as respiratory depression can progress to apnea without warning.

Documentation and Record Keeping

Accurate documentation of monitoring parameters is a professional obligation and a practical necessity in laboratory animal anesthesia. The anesthetic record should include baseline values, induction and maintenance agent doses, monitoring parameters at regular intervals, and any interventions performed. The interval between recordings depends on the stability of the patient and the invasiveness of the procedure. A minimum of every five minutes is standard for stable patients, with more frequent recording during induction, recovery, and periods of instability.

The record should note the monitoring method used for each parameter, as values obtained by different methods are not directly comparable. For example, a pulse oximetry heart rate may differ from an electrocardiographic rate during arrhythmias, and a Doppler pulse rate reflects perfusion instead of electrical activity. The record should also document the anesthetic depth assessment, including which reflexes were tested and the response obtained.

Poor reporting of anesthesia and monitoring in published studies is a recognized problem in laboratory animal research, with a systematic review of high-impact journals finding incomplete data on animal characteriztics and monitoring in many publications. Complete anesthetic records support both clinical care and the scientific validity of the research, as anesthetic agents influence physiologic parameters and can interfere with experimental results. Standardized conditions, including consistent monitoring and documentation, are necessary for the safe assessment of experimental outcomes.

Recognized Complications and Early Detection

Hypothermia remains the most frequent and consequential complication in small laboratory mammals under anesthesia. Mice lose heat rapidly through conduction, convection, radiation, and evaporative losses from the respiratory tract, and the resulting fall in body temperature slows drug metabolism, prolongs recovery, and depresses respiratory drive Gargiulo et al., mice anesthesia part I. Detect hypothermia early by continuous rectal or esophageal temperature measurement instead of intermittent checks, and respond when temperature falls below the species-specific baseline instead of waiting for a predetermined absolute threshold. A falling temperature accompanied by progressive bradycardia and reduced pulse quality suggests thermoregulatory failure has advanced to cardiovascular decompensation.

Respiratory depression is the second most common failure mode. Isoflurane produces a marked decrease in respiratory rate compared with injectable protocols, and this effect is dose-dependent Tsukamoto et al., vital signs monitoring in mice. Detect early by capnography or by direct observation of thoracic excursions, and distinguish a slow but adequate respiratory pattern from one that is both slow and shallow. A rising end-tidal carbon dioxide value with declining oxygen saturation indicates hypoventilation that requires immediate reduction of anesthetic delivery and, if necessary, assisted ventilation.

Bradycardia from injectable protocols deserves specific attention. Ketamine-xylazine combinations cause remarkable bradycardia in mice, whereas medetomidine-midazolam-butorphanol produces a relatively lower heart rate that reverses rapidly with atipamezole administration Tsukamoto et al., vital signs monitoring in mice. Detect early by continuous pulse oximetry or electrocardiography, and correlate the heart rate trend with the expected pharmacologic profile of the agent used. A heart rate that continues to fall despite stable anesthetic depth suggests drug effect instead of inadequate anesthesia.

Common Errors and Corrective Actions

Less experienced clinicians frequently misjudge anesthetic depth in small mammals because reflex responses differ from those in dogs and cats. The pedal withdrawal reflex may persist at surgical depth in some mouse strains, and relying on a single reflex invites either over-anesthesia or inadequate analgesia. Use a composite assessment that combines reflex responses, respiratory pattern, heart rate trend, and mucous membrane color, and document each parameter separately Gargiulo et al., mice anesthesia part II.

A second common error is failure to calibrate monitoring equipment for small patients. Pulse oximeter probes designed for larger species often read inaccurately on mouse or rat digits, and blood pressure cuffs must match the limb circumference. Verify that the signal quality indicator is stable before recording any value, and confirm abnormal readings with a second method before adjusting anesthetic depth.

A third error is neglecting to account for the physiologic effects of the experimental procedure itself. Hemorheological parameters change during ischemia-reperfusion, sepsis, and tissue trauma, and these changes can confound both monitoring values and experimental endpoints Nemeth et al., hemorheological variables in surgical pathophysiology research. Interpret monitoring data in the context of the surgical model, not as isolated vital signs.

Limitations of the Evidence

The evidence base for monitoring parameters in laboratory animals is uneven. A systematic review of publications in high-impact journals found poor quality and frequency of reporting on anesthesia, monitoring, and animal characteriztics, which limits the ability to compare protocols across studies Uhlig et al., systematic review of anesthesia reporting. Many published ranges derive from single strains, single sex, or single protocols, and extrapolation across species and strains is uncertain.

Expert opinion still differs on the value of advanced monitoring modalities. Near-infrared spectroscopy can track mitochondrial cytochrome oxidation during hemorrhagic shock, but gastric cytochrome oxidation may fail to recover despite systemic evidence of adequate resuscitation Rhee et al., near-infrared spectroscopy during hemorrhagic shock. Whether such regional monitoring should guide clinical decisions in routine laboratory anesthesia remains contested.

Escalation and Consultation

Escalation is warranted when physiologic parameters cannot be stabilized despite appropriate adjustment of anesthetic delivery, thermal support, and fluid administration. Persistent hypoxemia, progressive bradycardia, or loss of pulse quality despite corrective action requires immediate intervention and, where available, consultation with a laboratory animal veterinarian. Institutional animal care and use programs provide oversight and veterinary support for anesthetic complications National Research Council, Guide for the Care and Use of Laboratory Animals.

Regulatory reporting obligations vary by jurisdiction. Investigators should consult their institutional animal care and use committee and applicable national standards, including those published by the World Organization for Animal Health, to determine whether an anesthetic death or serious complication must be reported WOAH terrestrial animal health standards. Refinement of anesthetic protocols based on adverse events is an expectation of responsible animal use NC3Rs guidance on refinement.

Frequently Asked Questions

What Minimum Monitoring Is Acceptable When Full Physiologic Monitoring Equipment Is Unavailable?

When pulse oximetry, capnography, or noninvasive blood pressure devices are not available, monitoring must rely on direct clinical assessment. Rectal temperature measurement with a calibrated thermometer is non-negotiable, as hypothermia develops rapidly in small mammals and confounds anesthetic depth and recovery. Respiratory rate and pattern should be observed directly, with attention to thoracic versus abdominal effort. Heart rate and pulse quality require either a Doppler flow detector or manual palpation of the femoral or carotid pulse. Reflex assessment, including pedal withdrawal and palpebral responses, provides depth information but lags behind physiologic changes. Document all findings at five-minute intervals. The National Research Council Guide for the Care and Use of Laboratory Animals requires that monitoring be appropriate to the procedure and species, and that personnel be trained to recognize anesthetic complications.

How Should Monitoring Be Adapted for Animals Undergoing Imaging Procedures?

Imaging imposes specific constraints on monitoring. Magnetic resonance imaging prohibits ferromagnetic equipment, so MRI-compatible pulse oximetry probes and fiberoptic temperature sensors are required. Radiofrequency interference can disrupt electrocardiography and pulse oximetry signals. Positioning within the bore limits direct visual assessment, making remote monitoring essential. Physiological parameters change with positioning and immobilisation, and anesthetic protocols must be adjusted accordingly. The institutional review of anesthetic considerations in preclinical imaging studies notes that anesthesia must adapt to the imaging technique, procedure length, and study aim. Respiratory rate is often the most reliable indicator of depth during imaging, as heart rate may be affected by contrast agents and positioning. Pre- and post-imaging temperature checks are mandatory, as heat loss is accelerated in cold imaging suites.

What Are the Practical Limits of Pulse Oximetry in Mice and Neonatal Rodents?

Pulse oximetry in mice is technically challenging and frequently unreliable. The small tissue volume and rapid heart rate, often exceeding 500 beats per minute, exceed the processing limits of many standard probes. Motion artifact and peripheral vasoconstriction from hypothermia or anesthetic agents further degrade signal quality. The comparative study of vital signs during injectable and inhalant anesthesia in mice reported that isoflurane produced the most stable oxygen saturation readings among the protocols tested, while injectable combinations showed greater variability. When pulse oximetry fails, clinical assessment of mucous membrane color, capillary refill time, and respiratory effort becomes the primary oxygenation monitor. For neonatal rodents, pulse oximetry is rarely feasible, and monitoring relies on visible respiratory effort, mucous membrane color, and response to stimulation. Consider pulse oximetry readings as trend indicators instead of absolute values.

How Should Monitoring Data Be Reported in Publications?

The systematic review of anesthesia reporting in high-impact journals found poor quality and frequency of reporting on monitoring parameters in experimental studies. This omission undermines reproducibility and complicates interpretation of physiologic data. Publications should specify the anesthetic protocol, monitoring equipment, and the frequency of parameter recording. Report baseline values, intra-anesthetic ranges, and any deviations requiring intervention. Include the method of temperature support and the ambient temperature. When physiologic parameters are study endpoints, describe how anesthetic effects were distinguished from experimental effects. The review of hemorheological variables in surgical pathophysiology research emphasizes that standardized experimental conditions, including anesthesia and monitoring, are essential for valid comparison of results across studies.

When Should Monitoring Be Escalated to Invasive Techniques?

Noninvasive monitoring may be insufficient for prolonged procedures, cardiovascular studies, or animals with compromised physiologic reserve. Direct arterial blood pressure measurement via carotid or femoral artery catheterization provides continuous, accurate pressure data and enables arterial blood gas sampling. Central venous pressure monitoring guides fluid therapy in major surgical models. Invasive monitoring carries risks of hemorrhage, thrombosis, and infection, and requires surgical skill and additional anesthesia time. The decision to escalate should be based on the procedure's expected blood loss, the animal's baseline health, and the study's endpoints. For survival surgery, invasive catheterization should be removed and vessels repaired before recovery. The MSD Veterinary Manual provides species-specific guidance on vascular access techniques and their complications.

How Should a Supervisor or IACUC Be Informed of Monitoring Deficiencies?

Monitoring deficiencies should be reported promptly and factually, with emphasis on animal welfare and study validity. Document the specific parameter that could not be assessed, the equipment failure or limitation, and the corrective actions taken. If a monitoring device is unavailable or malfunctioning, state the alternative methods used and their limitations. The NC3Rs resources on refinement provide frameworks for assessing and improving procedural welfare. Institutional animal care and use committees expect transparency about monitoring failures, particularly in survival procedures. Propose a corrective plan, such as equipment repair, replacement, or protocol modification. If a monitoring deficiency resulted in an adverse outcome, describe the event chronologically and identify the point at which intervention occurred. This information supports institutional learning and prevents recurrence.

Related Clinical & Scientific Guides

• Refining IACUC Protocols to Minimize Animal Pain and Distress
• Health Monitoring Programs for Laboratory Animal Facilities
• Anesthetic Risk Assessment in Laboratory Animals: Preoperative Evaluation

References and Further Reading

• Mice anesthesia, analgesia, and care, Part I: anesthetic considerations in preclinical research.. 2012.
• Vital signs monitoring during injectable and inhalant anesthesia in mice.. 2015.
• Mice anesthesia, analgesia, and care, Part II: anesthetic considerations in preclinical imaging studies.. 2012.
• Anesthesia and Monitoring in Small Laboratory Mammals Used in Anesthesiology, Respiratory and Critical Care Research: A Systematic Review on the Current Reporting in Top-10 Impact Factor Ranked Journals.. 2015.
• Effects and influencing factors on hemorheological variables taken into consideration in surgical pathophysiology research.. 2018.
• Near-infrared spectroscopy: continuous measurement of cytochrome oxidation during hemorrhagic shock.. 1997.
• Guide for the Care and Use of Laboratory Animals, 8th Edition. National Academies Press, 2011.
• NC3Rs Resources on Replacement, Reduction and Refinement. NC3Rs.
• MSD Veterinary Manual, Professional Edition. MSD Veterinary Manual.

Related Articles

• Anesthesia Monitoring for Laboratory Rats: Parameters and Equipment
• Anesthesia for Laboratory Rabbits: Protocols and Monitoring
• Anesthetic Considerations for Pregnant Laboratory Animals
• Anesthesia Machine Safety Checks for Laboratory Animal Use
• Anesthesia Equipment for Small Laboratory Animals: Setup and Maintenance

This article is educational professional reference material for veterinary audiences. It is not a substitute for veterinary diagnosis, individual clinical judgment, current product labeling, or applicable regulatory requirements.